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DEVELOPMENTAL DYNAMICS 244:1215–1248, 2015
DOI: 10.1002/DVDY.24308
RESEARCH ARTICLE
Deployment of Regulatory Genes During Gastrulation
and Germ Layer Specification in a Model Spiralian
Mollusc Crepidula
a
Kimberly J. Perry,1 Deirdre C. Lyons,2 Marta Truchado-Garcia,3 Antje H. L. Fischer,4,5 Lily W. Helfrich,5
Kimberly B. Johansson,5,6 Julie C. Diamond,5 Cristina Grande,3 and Jonathan Q. Henry1*
1
University of Illinois, Department of Cell and Developmental Biology, Urbana, Illinois
Biology Department, Duke University, Durham, North Carolina
3
Departamento de Biologıa Molecular and Centro de Biologıa Molecular, “Severo Ochoa” (CSIC, Universidad Aut
onoma de Madrid), Madrid, Spain
4
Department of Metabolic Biochemistry, Ludwig-Maximilians-University, Munich, Germany
5
Marine Biological Laboratory, Woods Hole, Massachusetts
6
Department of Molecular and Cellular Biology, Harvard University, Cambridge, Massachusetts
DEVELOPMENTAL DYNAMICS
2
Background: During gastrulation, endoderm and mesoderm are specified from a bipotential precursor (endomesoderm) that is
argued to be homologous across bilaterians. Spiralians also generate mesoderm from ectodermal precursors (ectomesoderm),
which arises near the blastopore. While a conserved gene regulatory network controls specification of endomesoderm in deuterostomes and ecdysozoans, little is known about genes controlling specification or behavior of either source of spiralian
mesoderm or the digestive tract. Results: Using the mollusc Crepidula, we examined conserved regulatory factors and compared their expression to fate maps to score expression in the germ layers, blastopore lip, and digestive tract. Many genes
were expressed in both ecto- and endomesoderm, but only five were expressed in ectomesoderm exclusively. The latter may
contribute to epithelial-to-mesenchymal transition seen in ectomesoderm. Conclusions: We present the first comparison of
genes expressed during spiralian gastrulation in the context of high-resolution fate maps. We found variation of genes
expressed in the blastopore lip, mouth, and cells that will form the anus. Shared expression of many genes in both mesodermal sources suggests that components of the conserved endomesoderm program were either co-opted for ectomesoderm formation or that ecto- and endomesoderm are derived from a common mesodermal precursor that became subdivided into
C 2015 Wiley Periodicals, Inc.
distinct domains during evolution. Developmental Dynamics 244:1215–1248, 2015. V
Key words: Spiralia; gastrulation; blastopore; mesoderm; endoderm; EMT; mouth; foregut; hindgut; anus; Mollusca
Submitted 1 April 2015; First Decision 16 July 2015; Accepted 16 July 2015; Published online 21 July 2015
Introduction
Most bilaterian taxa have three principle germ layers: ectoderm,
endoderm, and mesoderm. The invention of mesoderm was intimately tied to the generation of the diverse morphologies of triploblasts (Perez-Pomares and Mu~
noz-Chapuli, 2002; Martindale,
2005; Martindale and Hejnol, 2009). Much effort has centered on
deciphering the molecular basis for specifying distinct ectodermal, endodermal, and mesodermal germ layers during development and great progress has been made to understand germ layer
specification in deuterostomes (e.g., echinoderms, chordates),
ecdysozoans (e.g., fly, nematode), and cnidarians (Byrum and
Martindale, 2004; Martindale et al., 2004; Magie et al., 2007;
Sawyer et al., 2010; Gorfinkiel et al., 2011; Peter and Davidson,
2011; R€
ottinger et al., 2012; Solnica-Krezel and Sepich, 2012). In
Kimberly J. Perry and Deirdre C Lyons are co-first authors.
*Correspondence to: Jonathan Q. Henry, Department of Cell and Developmental Biology, University of Illinois, 601 S. Goodwin Avenue, Urban, IL
61801. E-mail: [email protected]
contrast, germ layer specification is not well understood in the
Spiralia, a major branch of bilaterians (Boyle et al., 2014; Passamaneck et al., 2015). The Spiralia contains animals such as
annelids, molluscs, brachiopods, phoronids, rotifers, among
others, with diverse larval and adult body plans (Hejnol, 2010;
Henry, 2014). Many members of the Spiralia exhibit a highly
conserved spiral cleavage pattern and cell lineage fate map (Hejnol, 2010; Lambert, 2010; Henry, 2014). This characteristic spiral
cleavage program, together with molecular phylogenetic analyses, reveals the close evolutionary relationships amongst members of the Spiralia (Boyer et al., 1996; Henry and Martindale,
1999; Dunn et al., 2008; Giribet, 2008; Hejnol, 2010; Lambert,
2010; Henry, 2014). Thus understanding germ layer specification
in the Spiralia, particularly in those with spiral cleavage, which is
believed to be the ancestral condition for this clade, should
expand our understanding of how this clade evolved.
Article is online at: http://onlinelibrary.wiley.com/doi/10.1002/dvdy.
24308/abstract
C 2015 Wiley Periodicals, Inc.
V
1215
DEVELOPMENTAL DYNAMICS
1216 PERRY ET AL.
Ectoderm, endoderm, and mesoderm are segregated during the
process of gastrulation, which occurs as the result of cellular signaling, and morphogenetic rearrangement (Stern, 2004). Morphogenetic events associated with the process of gastrulation and
germ layer formation also have a direct bearing on the formation
of the digestive tract, including structures such as the mouth and
the anus (Hejnol and Martindale, 2008; Lyons and Henry, 2014).
Cell lineage and fate mapping studies show that mesoderm and
endoderm usually arise from common endomesoderm progenitor
cells, and that this condition appears to be conserved across metazoans, including in ctenophores, acoels, ecdysozoans, spiralians,
echinoderms, urochordates, and vertebrates (Nishida and Satoh,
1985; Martindale and Henry, 1999; Logan et al., 1999; Henry
et al., 2000; Kimelman and Griffin, 2000; Fukuda and Kikuchi,
2005; Maduro, 2006; Gilles et al., 2007; Ryan et al., 2013). Mesoderm likely evolved from endoderm, since many genes associated
with bilaterian mesoderm cell types are expressed within the
endoderm of basally branching animals like cnidarians (Martindale et al., 2004; R€ottinger et al., 2012). It is now common to
build gene regulatory networks (GRNs) for specification of endoderm, and mesoderm in order to understand how endoderm and
mesoderm become distinct cell types during development and
evolution (Hinman et al., 2003; Maduro, 2006; Peter and Davidson, 2011; R€
ottinger et al, 2012). Such studies have revealed
highly conserved core regulatory components and wiring that are
involved in endomesoderm specification and diversification.
Comparing GRNs across species can reveal which sub-circuits of
the network are most conserved, and how they have diverged
over evolutionary time.
The most complete endomesoderm GRN is that of the sea urchin
(Oliveri et al., 2008; Peter and Davidson, 2011; http://sugp.caltech.edu/endomes). It often serves as a framework for examining
germ layer specification in other organisms. Additional GRNs
have been reconstructed for sea stars (Hinman and Davidson,
2003; Hinman et al., 2003; McCauley et al., 2015), vertebrates
(Chan et al., 2009), nematode (Maduro and Rothman, 2002;
Maduro, 2006; Owraghi et al., 2010), and sea anemone (R€
ottinger
et al., 2012). To date, no endomesoderm GRN has been built for a
spiralian. A necessary pre-requisite for generating a spiralian
GRN requires that three types of data must be acquired: (1) fate
mapping data showing when and from where ectoderm, endoderm, and mesoderm cell types arise, (2) experimental perturbations to establish to what extent specification is accomplished cell
autonomously, versus as a result of inter-cellular signaling, and
(3) gene expression data showing where germ layer specification
and differentiation genes are expressed, relative to the fate map.
The highly stereotyped early spiral cleavage program has made
it possible to generate high-quality fate maps using modern intracellular lineage tracing techniques in several species (Boyer et al.,
1996; Dictus and Damen, 1997; Henry and Martindale, 1998;
Goto et al., 1999; Hejnol et al., 2007; Meyer et al., 2010a,b; Gline
et al., 2011; Chan and Lambert, 2014). These lineage studies have
revealed an interesting fact: unlike many other animal groups,
spiralians have two sources of mesoderm (Lillie, 1895; Wilson,
1898; Henry and Martindale, 1999; Henry et al., 2000; Martindale, 2005; Lyons and Henry, 2014). Endomesoderm typically
arises from the 4d micromere, and gives rise to heart, muscle, kidney, germ line, and additional mesenchyme (Lambert, 2008;
Goulding, 2009; Gline et al., 2011; Lyons et al., 2012; Fischer and
Arendt, 2013; Lyons and Henry, 2014), suggesting that this source
of spiralian mesoderm is homologous to mesoderm in other bilat-
erians. In the snail Crepidula, the 4d cell divides bilaterally, producing two stem cells (teloblasts) that divide synchronously and
give rise to daughter cells that are strictly endodermal or mesodermal; cell ablation experiments demonstrated that these fates are
determined autonomously (Lyons et al., 2012). Here when we refer
to expression in endomesoderm, we mean specifically the mesodermal derivatives of 4d. However, spiralian mesoderm is also
derived from ectomesoderm, which typically gives rise to larval
tissues (e.g., muscles), as well as some adult tissues (Boyer et al.,
1996; Lyons and Henry, 2014; Chan and Lambert, 2014). The cells
that give rise to ectomesoderm vary between species, but in many
spiralians ectomesoderm arises primarily from 3a and 3b micromeres, which is argued to be the ancestral condition (Dictus and
Damen, 1997; Byrum and Martindale, 2004; Hejnol et al., 2007;
Chan and Lambert, 2014; Lyons and Henry, 2014). At a molecular
level, we do not know whether these two types of mesodermal
precursors are regulated by the same or different specification
programs, but some investigators have suggested that spiralian
endomesoderm is similar to endomesoderm in other metazoans
on the basis of shared expression of regulatory genes (Lartillot
et al., 2002b; Arenas-Mena, 2006, 2013; Dill et al., 2007; Henry
et al., 2008; Pfeifer et al., 2013; Boyle et al., 2014).
While many spiralian fate maps have traced later development
of larval structures, few fate maps have traced the behavior of cells
during gastrulation, when germ layers typically first become specified. Recently we completed such fate maps for the snail, Crepidula
fornicata (Lyons et al., 2012, 2015), making it possible to examine
gene expression patterns in the context of the highly stereotyped
spiralian fate map and germ layer formation in a mollusc. In the
present study we cloned fifteen genes from C. fornicata that are
known to be involved in gastrulation, endomesoderm specification,
and gut development in other model systems. We analyzed their
expression patterns in order to compare the lineage-specific origins
of the germ layers, mouth, foregut, and hindgut, with what is currently known in other metazoans. For many of the genes examined,
we showed expression in both endo- and ectomesoderm. This result
suggests that in Spiralia, either a conserved metazoan endomesoderm GRN (Davidson et al., 2002; Loose and Patient, 2004; Davidson and Erwin, 2006; Peter and Davidson, 2011) was co-opted for
ectomesoderm specification, or alternatively, that ecto- and endomesoderm were derived from common mesodermal precursors that
became segregated into distinct domains during the subsequent
course of evolution.
We also examined gene expression for common markers of the
digestive system, including genes known to be involved in mouth
and hindgut specification. Expression as it relates to the embryonic
blastopore is presented for various stages of gastrulation and early
organogenesis, and is further compared to other members of the
Spiralia and Bilateria in order to compare the diverse developmental origins of these structures. We found shared expression in both
the ectodermal foregut and the endodermal hindgut, which may
reveal common aspects in their specification, as well.
Results
Overview of Cleavage, Gastrulation and
Organogenesis in Crepidula
Spiral cleavage involves alternating sets of oblique cell divisions
that form staggered quartets of micromeres at the animal pole of
the embryo. The first two cell divisions generate four blastomeres
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1217
occupying each of the basic embryonic quadrants (termed A, B,
C, and D; see Conklin, 1897). Each of these cells subsequently
generates a series of typically smaller animal daughter cells
termed “micromeres,” which are formed in alternating clockwise
and counterclockwise orientations around the animal-vegetal
axis, whereas the four vegetal-most cells are typically larger and
termed “macromeres.” Animal micromeres are designated with
lowercase letters, while the vegetal macromeres are designated
with uppercase letters. Hence, the first quartet of micromeres is
named 1a, 1b, 1c, and 1d, while the corresponding macromeres
are named 1A, 1B, 1C, and 1D. The second quartet is named 2a,
2b, 2c, and 2d, and the corresponding macromeres are named 2A,
2B, 2C, and 2D, and so on.
The process of gastrulation follows early cleavage events and
has recently been examined in depth in C. fornicata using in vivo
imaging and cell lineage analyses (Lyons et al., 2015). Gastrulation occurs by epiboly, and is characterized by expansion of the
progeny of the first through third quartet animal micromeres
toward the equatorial zone and ultimately the vegetal pole
(approximately 48–117 hpf; all staging follows that established
in Lyons and Henry, 2014, and Lyons et al., 2015). During this
process, animal cap cells surround vegetal endodermal precursors
(the fourth quartet micromeres: 4a–4c; and the macromeres: 4A–
4D). As epiboly progresses, the leading edge of animal micromeres (i.e., the blastopore lip) begin to spread and cover these
underlying cells (round stages, 48–117hpf). Eventually, the circumference of the blastopore lip constricts as it approaches the
vegetal pole of the embryo, and the cells around the lip (derived
from progeny of the second and third quartet of micromeres; 2a–
2d and 3a–3d; Fig. 1) undergo rearrangement around the open
blastopore. The blastopore, as it relates to C. fornicata, is defined
as the site where presumptive endoderm and mesoderm are internalized during gastrulation. In C. fornicata, this opening never
fully closes and ultimately gives rise to the mouth and ectodermal
foregut (or esophagus). Cells of the posterior lip of the blastopore
(progeny of 2d) also contribute to the anus (Lyons and Henry,
2014; Lyons et al., 2015). Specifically, during later stages of epiboly (beginning at 97 hpf), the posterior lip of the blastopore closes
by a zippering process that involves convergence and extension
(CE; Lyons et al., 2015). At this time, the cells that are ultimately
involved in the formation of the anus are displaced towards the
posterior pole of the embryo, away from the persistent opening of
the blastopore (described further below).
Mesoderm in C. fornicata is derived from two sources: endomesoderm (derived from one of the fourth quartet micromeres,
4d; Fig. 1), and ectomesoderm (derived from ectodermal bipotential progeny of 3a and 3b at the very anterior-lateral edges of the
blastopore; Fig. 1). During epiboly, progeny of 4d reside at a posterior location adjacent to the blastopore lip (Lyons et al., 2015).
For the expression data represented in the next section, we refer
to any expression (after 90 hpf) in the 4d lineage as either being
expressed in the mesodermal derivatives as “endomesoderm,” or
in the 4d-derived endodermal progeny, which contributes to the
“hindgut precursors” or “hindgut rudiment.” Following blastopore constriction, the embryo undergoes anterior-posterior axial
elongation (elongation stages, 117–170hpf) and this axial elongation results in the mouth becoming displaced towards the anterior pole of the embryo (Fig. 1; Lyons and Henry, 2014). By 145
hpf (later stages of elongation just prior to organogenesis stages),
the morphogenetic events that make the mouth and esophagus
are largely completed, and subsequent morphogenesis results in
axial elongation of the embryo. Posterior closure of the blastopore lip and axial elongation displaces the ectodermal progeny of
2d (including the 2d2 derived anus anlagen), the terminal cells
(3c2/3d2-derivatives), and the endodermal hindgut, away from
the mouth.
During organogenesis stages, the ciliated prototroch forms perpendicular to the anterior-posterior axis and separates the ectoderm of the developing larvae into different regions. The anterior
region consists of pre-trochal ectoderm (derived from the first
quartet and includes anterior ectoderm of the velum, apical
organ, and apical ganglia, 1q; Fig. 1). The post-trochal region
that lies posterior to the mouth consists of ectoderm derived from
second and third quartet micromeres (2q, 3a; Fig. 1). During later
stages of organogenesis (196hpf), the rudiments of the velar lobes
become more pronounced and the ciliated apical plate shifts ventrally within the developing head. Prominent ciliated bands
involved in larval feeding and locomotion form at the edge of the
developing velum, and include the food groove, and primary and
secondary ciliary bands (which Hejnol et al., 2007, describe as the
prototroch and metatroch, respectively). In the post-trochal
region, the shell gland forms on the dorsal surface, the foot rudiment and operculum develop on the ventral surface, and the
paired bilateral larval external kidneys or absorptive cells develop
on the left and right lateral sides (Rivest, 1992; Lyons et al.,
2012). The hindgut rudiment also forms within the post-trochal
region along the posterior ventral midline, but as development
progresses this structure shifts towards the right lateral side (see
Hejnol et al., 2007; Henry et al., 2010a; Lyons et al., 2012, 2015).
The veliger larval stage is reached by 10 days of development.
The anus, which will be located at the distal tip of the hindgut,
does not open until later, at approximately 12 days of development (veliger larval stage; Lyons et al., 2015).
In situ Expression Patterns
Endomesoderm
Beta-catenin (ctnnb). Beta-catenin is a multifunctional protein
involved in cell adhesion, cell signaling, and transcriptional regulation. It is broadly expressed, given its roles in cell adhesion. It
has also been shown to play conserved roles during gastrulation
and endomesoderm specification, and serves as a key upstream
component of the sea urchin endomesoderm GRN (Wikramanayake et al., 1998, 2003; Logan et al., 1999; Imai et al., 2000;
Miyawaki et al., 2003; Kawai et al., 2007; R€
ottinger et al., 2012).
Active ctnnb signaling occurs with the nuclearization of this protein that serves as an intracellular signal transducer to regulate
downstream targets, such as those involved in the canonical Wnt
signaling pathway. The expression of ctnnb has been described
for C. fornicata (Henry et al., 2010), but that study did not specifically describe expression during gastrulation stages. Here we
report on the mRNA expression patterns, which do not necessarily indicate active ctnnb signaling.
The expression pattern of ctnnb at early stages of development
is dynamic (Figs. 1 and 2A-F; Table 1; J. Q. Henry et al., 2010). In
a previous study we showed that expression is seen weakly in
macromeres and animal micromeres (quadrants A–D), and
depending on the cell cycle, ctnnb becomes more highly localized
to a region adjacent to the nuclei, which we presume to be the
centrosomes (Henry et al., 2010c). By mid to late gastrulation
more intensely labeled cells are observed around the entire edge
DEVELOPMENTAL DYNAMICS
1218 PERRY ET AL.
Fig. 1. Lineage and expression summary during development in C. fornicata. Fate map for various stages of C. fornicata development is represented along the top row with groups of clones highlighted according to their clonal lineage in various colors. Key is located at the bottom right
corner of the figure. Illustrations are shown as vegetal/ventral views with anterior to the top of the figure. Below is a summary of each gene
expression pattern presented for three stages of development representing epiboly (94hpf), elongation (120–130hpf), and late elongation/early
organogenesis (140–145hpf). If no pattern is displayed, then no expression was observed at that particular stage. The blastopore (bp) is outlined
for each stage with a dashed grey line. “em” represents ectomesoderm.
of the blastopore (Fig. 2E–I). Signal is noted particularly along
the posterior edge of the blastopore lip, along with cells exhibiting higher levels of expression extending from the posterior blastopore along the ventral midline, towards the more posterior
regions (ectodermal cells derived form 3c and 3d, Fig. 2G–H).
Fainter label also becomes visible in some posterior progeny,
lateral to the ventral midline. During later elongation stages,
expression is observed in the anterior ectoderm derived from the
1st quartet micromeres and in posterior ectoderm (mainly progeny
of 2d) in a radiating pattern that extends from the blastopore and
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1219
Fig. 2. Expression of ctnnb during early development in C. fornicata. A–T: ctnnb expression in embryos ranging from 80–170hpf (early epiboly–early
organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each panel
(an., animal; veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Animal view of ctnnb
expression (80hpf) (A) and corresponding DAPI labeled nuclei (B). Embryonic quadrants A–D are labeled within each panel of A–D. Vegetal view of same
80hpf specimen in A,B as epiboly begins, showing the localization of ctnnb expression (C) and corresponding DAPI-labeled nuclei (D). Flattened round
stage embryo (E) and corresponding DAPI image (F), and early elongation embryo with expression around the blastopore (G) with corresponding DAPI
labeling (H). Expression during elongation is represented in I with corresponding DAPI labeling in J. A dorsal view of expression from the same embryo
show in I–J is shown in (K) and corresponding DAPI labeled nuclei (L). Various views of expression are shown from the same late elongating ovoid specimen in M–P, and similarly the same organogenesis specimen in Q–T. Structures are labeled as follows: bp, blastopore; ec, ectomesoderm; hr, hindgut
rudiment; mo, mouth; sg, shell gland; st, stomodeum. Scale bar in T ¼ 50 mm.
1220 PERRY ET AL.
DEVELOPMENTAL DYNAMICS
TABLE 1. Grids Detailing Expression in Specific Tissues/Rudiments During Embryogenesis in C. fornicata (3 Representative Time Periods)a
a
Genes are listed along the top of the grids and regions of expression are noted along the left side of the grids. The embryonic
range of development is noted at the top left of each expression grid. Shaded boxes represent areas where expression was noted
and un-shaded boxes represent areas where no expression was detected. The embryonic range of development examined is noted
at the top left of each expression grid, including epiboly (90–117hpf), elongation (120–137hpf) and late stages of elongation/early
organogenesis (140–170hpf).
developing mouth (Fig. 2I–L). Expression is somewhat diminished
by early organogenesis stages, but broadens during later organogenesis stages and is observed around the mouth, the shell gland,
and a proliferative zone located on the left-and right-lateral
post-trochal area (Fig. 2M–T). Pre-larval expression also appears
to be somewhat more intense on the right side.
Expression is also seen in the progeny of 4d. ctnnb expression
was observed in organogenesis specimens, where it is noted
more strongly in progeny of 4d, particularly the hindgut rudiment (Fig. 2Q–S).
Orthodenticle (otx). Orthodenticle is a homeobox transcription
factor with conserved roles in the specification of anterior structures, including those of the CNS and the stomodeum (Cohen and
J€
urgens, 1990; Finkelstein et al., 1990; Acampora et al., 2000;
Boyle et al., 2014).
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1221
Fig. 3. Expression of otx during early development in C. fornicata. A–P: otx expression in embryos ranging from 99–165hpf (late epiboly through
elongation stages). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each
panel (veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Mid-epiboly embryo
(A) and corresponding DAPI image (B), and later epiboly embryo with expression around the blastopore (C) with corresponding DAPI labeling (D).
Expression during elongation is represented in E with corresponding DAPI labeling in F. Ventral expression in an elongating oval stage is shown in
G with corresponding dorsal view in H. Various views of expression are shown from the same late ovoid specimen in I–L, and similarly the same
organogenesis specimen in M–P. Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in I, J, L–N, P. Structures are
labeled following designations used in Figure 2 and as follows: fr, foot rudiment; np, neural precursors; vr, velar rudiment. Scale bar in P ¼ 50mm.
otx does not exhibit a distinct localized expression pattern during early cleavage stages, but there appears to be faint widespread expression throughout the embryo. During early to midgastrula stages, more pronounced expression is seen around the
blastopore (Figs. 1, 3A–D; Table 1), and later gastrula embryos
have expression around the entire blastopore (Fig. 3E–F). Mid to
late gastrula embryos display otx labeled cells along the ventral
midline posterior to the blastopore (Fig. 3C–F). During organo-
genesis, expression is noted in the anterior mouth and esophagus,
with less intense expression along the posterior edge of the
mouth and foregut (Fig. 3G, I–K, M–O). otx expression is widespread in the ectoderm at later stages of development and noted
asymmetrically in a patch of cells on the right side of the embryo
in the post-trochal region (Fig. 3I–J, L–N, P). Expression is also
observed anterior to the mouth in head ectoderm and scattered
cells likely to be the neuronal precursors (Fig. 3G, H, J, M–O). The
DEVELOPMENTAL DYNAMICS
1222 PERRY ET AL.
Fig. 4. Expression of foxA during early development in C. fornicata. A–T: foxA expression in embryos ranging from 24-cell-170hpf (early cleavage to
early stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each panel (an., animal; veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images.
Animal view of foxA expression in cleavage embryo (A) and corresponding DAPI labeled nuclei (B). Embryonic quadrants A–D are labeled within each
panel of A–D. Animal view of foxA expression in 45hpf specimen (C) and corresponding DAPI-labeled nuclei (D). A dorsal view of a 48hpf specimen
(E) and corresponding DAPI image (F) showing expression that is localized to the teloblasts progeny (1mL, 1mR, 3mL, 3mR). Flat round embryo with
expression at the posterior region of the blastopore (G) with corresponding DAPI labeling (H). foxA expression during an epiboly stage (I) with corresponding DAPI labeling (J), and an ovoid stage (K) with corresponding DAPI labeling (L). Various views of expression are shown from the same late
ovoid specimen in M–P, and similarly the same organogenesis specimen in Q–T. Black arrowheads point to asymmetric post-trochal ectodermal
patch of expression in Q, T. Structures are labeled following designations used in Figures 2 and 3. Scale bar in T ¼ 50 mm.
DEVELOPMENTAL DYNAMICS
Fig. 5. Expression of bra during early development in C. fornicata. A–X: bra expression in embryos ranging from 8-cell–170hpf (early cleavage to
early stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right
corner of each panel (an., animal; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Animal view of bra
expression in early cleavage embryo (A) and corresponding DAPI-labeled nuclei (B). Embryonic quadrants A–D are labeled within each panel of A–
F. Animal view of 48hpf specimen (C) and corresponding DAPI-labeled nuclei (D) showing bra expression in the teloblasts (3ML, 3MR) along with
the dorsal view of the same embryo (E) and corresponding DAPI labeled nuclei (F). Early oval embryo with expression at the posterior region of
the blastopore (G) with corresponding DAPI labeling (H). bra expression is shown in an early ovoid stage (I). A later ovoid stage (J) is shown with
corresponding DAPI labeling (K), as well as anti-acetylated tubulin labeling (L, green fluorescence). A posterior view of the same specimen from J–
L is shown (M) with the corresponding DAPI labeling (N). A late oval stage (elongated) is shown (O) as well as a similarly staged specimen showing
anti-acetylated tubulin labeling (P, green fluorescence). Various views of expression are shown from the same late ovoid specimen in Q–T, and similarly the same organogenesis specimen in U–X. Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in U, V, and
X. Structures are labeled following designations used in Figures 2 and 3, and as follows: ap, apical plate; nt, neurotroch; tb, teloblasts; tc, terminal
cells; vm, ventral midline. Scale bar in X ¼ 50 mm.
DEVELOPMENTAL DYNAMICS
1224 PERRY ET AL.
Fig. 6. Expression of cdx during early development in C. fornicata. A–X: cdx expression in embryos ranging from 16-cell–170hpf (early cleavage to early
stages of organogenesis). Orientation of specimens is located in the bottom right corner of each panel (an., animal; veg., vegetal; veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Animal view of cdx expression in early cleavage embryo
(A) and corresponding DAPI labeled nuclei (B), as well as more advanced cleavage embryo (C) and corresponding DAPI nuclei (D). Embryonic quadrants
A–D are labeled within each panel of A–F. Vegetal view of 80hpf specimen (E) and corresponding DAPI labeled nuclei (F). Epiboly specimen is shown with
blastopore expression (G) and corresponding DAPI labeling (H). cdx expression is shown at an ovoid stage (I) with corresponding DAPI labeling (J), as
well as a posterior view of the same embryo (K) with corresponding DAPI labeling (L). An elongating stage is shown (M) with corresponding DAPI labeling
(N), as well as a slightly older ovoid stage (O) and DAPI labeling (P). Various views of expression are shown from a single late ovoid specimen in Q–T, and
also a single organogenesis stage specimen in U-X. Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in Q, R, T–V, and
X. Structures are labeled following designations used in Figures 2, 3, and 5. Scale bar in X ¼ 50 mm.
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1225
DEVELOPMENTAL DYNAMICS
developing velum expresses otx (Fig. 3G, I–K, M–O), as do cells in
the foot rudiment (Fig. 3N) and in the proliferative zone of the
shell gland (Fig. 3I, K, L).
otx can also be observed in ectomesodermal derivatives (3a2,
3b2; Fig. 3C–G), as well as the ectomesoderm cells undergoing
EMT. Expression is also observed in both the endomesodermal
progeny of the mesentoblast (2mL and 2mR) and in the endodermal hindgut rudiment (Fig. 3E–G, I–K, M–O). Relatively intense
expression is present in the teloblast cells at certain stages.
Forkhead box A (foxA). FoxA is a member of the forkhead
box, helix-turn-helix class of transcription factors (Kaufmann
and Knochel, 1996). Fox genes are known to play roles in specification and differentiation of endodermal structures and are
thought to regulate the fates of both ectomesoderm (ArenasMena, 2006) and endomesoderm (Olsen and Jeffrey, 1997; Martindale et al., 2004; Oliveri et al, 2006; Boyle and Seaver, 2008,
2010).
During early cleavage stages in C. fornicata, foxA mRNA is
localized to regions adjacent to the nuclei in the A–D macromeres, which we presume to be the centrosomes (Figs. 1, 4A–D).
Expression is seen during epiboly in cells of the blastopore lip
(Fig. 4G–J; Table 1), and in particular cells of the posterior lip,
which undergo convergent extension (Fig. 4I–J). As development
continues, expression is seen in all cells of the developing mouth
and esophagus, being somewhat more highly expressed in the
anterior regions of both structures (Fig. 4K–L). During organogenesis, expression appears in cells posterior to and along the rim
of the velum (Fig. 4M–S). Expression is also seen in the foot rudiment and the shell gland (Fig. 4M, O–T).
Expression is detected in progeny of the mesentoblast (4d),
including progenitors of the hindgut during early to mid stages
of gastrulation (e.g, 1mL, 1mR, 3mL, 3mR; Fig. 4E, F). Expression
is also apparent in the progenitors of the ectomesoderm (progeny
of 3a2, 3b2; Fig. 4K, L), and in scattered mesenchymal progeny of
3a2 and 3b2. During stages of organogenesis, expression appears
to be lost in ectomesoderm that has migrated away from the
region of the mouth, though expression continues in the endoderm of the developing hindgut rudiment (Fig. 4M–O; Q–S). foxA
is asymmetrically expressed in post-trochal ectoderm with more
expression in a patch on the right side. (Fig. 4Q, T). Similar to C.
fornicata, foxA is also expressed asymmetrically in one blastomere on the right side of the post-trochal region in Patella (possibly the ectodermal midline stem cell; Lartillot et al., 2002a).
Brachyury (bra). Brachyury is a T-box transcription factor that
plays a key role in specifying cells of the blastopore (Shoguchi
et al., 1999; Arendt et al., 2001; Technau, 2001; Gross and
McClay, 2002), regulating genes involved in gastrulation-specific
morphogenetic movements (i.e., convergent extension; Arendt,
2004), and the development and specification of the adult mesoderm (Peterson et al., 1999; Technau, 2001; Gross and McClay,
2002; Boyle et al., 2014). It also regulates the development of the
anterior-posterior axis (Arendt et al., 2001; Lartillot et al., 2002b;
Koop et al., 2007). More recently a study in the cnidarian Acropora suggests a role for bra in demarcating ectoderm from endoderm (Hayward et al., 2015).
In C. fornicata, bra expression appears to be dynamic and
mainly restricted to the vegetal macromeres during early cleavage
stages, where it localizes to regions adjacent to the nuclei during
interphase (presumably the centrosomes, Figs. 1, 5A, B; Table 1).
Centrosomal localization has been reported previously in the
molluscs, Ilyanassa (Lambert and Nagy, 2002; Kingsley et al.,
2007) and Crepidula (Henry et al., 2010). After the 24-cell stage,
localization appears mainly in cells of the D-quadrant (Fig. 5C–
F). Localization is observed during early gastrulation around the
lip of the entire blastopore with more concentrated expression
seen along the posterior edge (Fig. 5G, H) and in ectoderm posterior to the blastopore lip. This expression pattern is reminiscent
of that seen for ctnnb. There is localization of bra in some cells
derived from 3c1 and 3d1 to the sides of the ventral midline (Fig.
5I). Signal is also observed in progeny of 3a2 and 3b2, which give
rise to ectomesoderm, and are located on the lateral sides of the
blastopore. Unlike in Capitella, we did not observe asymmetric
bra expression to the left side of the blastopore (Boyle et al.,
2014). Localization of bra in gastrula stage embryos is also noted
in what are presumed to be neuronal precursors from scattered
anterior 1q progeny. At later stages, expression is seen in the cells
that undergo convergent extension (from 3c2 and 3d2), which
originally occupied the posterior blastopore lip (including the terminal cells; Fig. 5O–P; Lyons et al., 2015), and come to reside
along the ventral midline (Fig. 5I, O). During later gastrula stages
expression continues in cells around the entire blastopore lip.
During cleavage stages, two areas of expression are also
observed in the mesodermal left and right teloblasts derived from
4d (Fig. 5J–N), and at later stages of gastrulation, prominent
expression continues in the teloblasts and in endomesodermal
progeny. Pre-larval expression appears around the entire mouth,
in the esophagus, in both mesodermal and endodermal progeny
of 4d, including the hindgut rudiment, in intense spots associated
with nuclei in some cells of the anterior/apical plate ectoderm, in
the prototroch cells (derived from 1q), and in the more ventrallateral progeny of 2b1 (neuronal cells) and 2b2 (Fig. 5Q–X). Proximal lateral cells of the foot rudiment also express bra (derived
from 3c1 and 3d1, Fig. 5U–W).
Several bra in situ specimens were also labeled with acetylated
tubulin antibody, which labels ciliated cells that appear during
later stages of gastrulation (during elongation) and organogenesis
stages. These ciliated cells are found in ectodermal regions of the
mouth, apical plate, and cells located posterior to the mouth
along the ventral midline including the two terminal cells (Lyons
et al., 2015; Fig. 5L). Acetylated tubulin labeling corresponds
with regions along the ventral midline where bra expression is
also noted. During later elongation stages and organogenesis, the
two ciliated terminal cells have migrated towards the posterior
pole (Fig. 5P), and lie in a region where bra mRNA is also
observed (Fig. 5O, P).
Digestive Tract Markers
Caudal (cdx). Caudal is a hox-related homeobox transcription
factor important during early embryonic development and in the
process of gastrulation. At later stages, cdx is involved in the
specification of posterior fates, including the hindgut (Schulz and
Tautz, 1995; Wu and Lengyel, 1998; Moreno and Morata, 1999;
Lartillot et al., 2002b).
During cleavage stages, cdx is localized in specific cells of all
four quadrants of the embryo and is particularly localized to
regions adjacent to the nuclei, which we presume to be the centrosomes of the macromeres, and to some extent also in the
micromeres (Figs. 1, 6A–F; Table 1). At later stages expression
appears to be more widespread and is seen in particular around
DEVELOPMENTAL DYNAMICS
1226 PERRY ET AL.
Fig. 7. Expression of gsc during early development in C. fornicata. A–T: gsc expression in embryos ranging from 16-cell–145hpf (early cleavage
to later stages of elongation). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each panel (an., animal; veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield
images. Animal view of gsc expression in 16-cell cleavage stage (A) and corresponding DAPI-labeled nuclei (B), as well as a 27-cell cleavage
stage (C) and corresponding DAPI nuclei (D). Embryonic quadrants A–D are labeled within each panel of A–F. Animal view of late cleavage specimen (E), and corresponding DAPI-labeled nuclei (F). Flattened round stage embryo (G) and corresponding DAPI image (H), and late epiboly
embryo with expression around the blastopore (I) with corresponding DAPI labeling (J). The embryo shown in I, J is represented in a posterior
view (K) with corresponding DAPI labeling (L). Expression during elongation is represented in (M) with corresponding DAPI labeling in (N) and in a
later specimen (O) with DAPI labeling (P). Various views of expression are shown from the same late ovoid specimen in Q–T. Black arrowheads
point to asymmetric post-trochal ectodermal patch of expression in Q, R, and T. Structures are labeled following designations used in Figures 2
and 5. Scale bar in T ¼ 50 mm.
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1227
Fig. 8. Expression of hex during early development in C. fornicata. A–H: hex expression in embryos ranging from 110–196hpf (late epiboly to late
stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner
of each panel (veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany one brightfield image. Vegetal ventral
view of hex expression very faintly along the anterior region of the blastopore (A) and corresponding DAPI labeled nuclei (B). Ventral view of hex
expression in ovoid stage specimen (C) and even later ovoid stage (D). Various views of the same specimen during organogensis are represented
in (E-H). Structures are labeled following designations used in Figures 2 and 3. Scale bar in H ¼ 50 mm.
posterior and anterior-lateral cells of the blastopore (Fig. 6G,H).
Though between 120–137hpf (elongating stage), it is excluded
from the very anterior region of the blastopore lip. cdx is
observed in posterior ectoderm, and possibly in some cells that
are located closest to the ventral midline (Fig. 6I–P). Expression is
widespread and includes other ectodermal cells, including an
asymmetric patch in the right post-trochal region (Fig. 6I–L). At
later stages, cdx is seen in the cells that undergo convergent
extension, which originally comprised the posterior blastopore
lip. This includes the two terminal cells, but expression at later
stages appears to be extinguished in these cells (Fig. 6K, L, O, P).
As development continues, cdx is expressed in all cells of the
developing mouth and the esophagus. Pre-larval expression is
seen in the cells of the hindgut, shell gland, velar lobes, and foot
rudiment (Fig. 6Q, X). Noticeably, cdx is asymmetrically
expressed in the ectoderm with more expression in a patch on the
right side of the post-trochal region as compared with the left
side. Asymmetric expression of cdx was also noted in a similar
region in Patella (Lartillot et al., 2002b).
Expression is also seen in progeny of the mesentoblast (4d),
including distinct expression in the two teloblasts at certain
stages (Fig. 6K–N). At later stages, prominent expression is seen
in both endomesoderm and the developing hindgut rudiment
(Fig. 6R, U, V). Furthermore, expression is apparent in the progenitors of ectomesoderm (Fig. 6Q–X), but is not apparent once
the progeny have undergone EMT.
Goosecoid (gsc). Goosecoid is a homeobox transcription factor
expressed during gastrulation. It is expressed during the development of mesendodermal fates in many metazoans, and promotes
the development of the dorsal organizer in chordates (Angerer
et al., 2001; Lartillot et al., 2002a; Boyle and Seaver, 2008).
During early cleavage stages gsc mRNA is concentrated in
regions adjacent to the nuclei, which we presume to be the centrosomes of the A–D macromeres, but is also expressed in animal
micromeres (Figs. 1, 7A–F; Table 1). Faint expression is widespread throughout the embryo, but during epiboly it is more concentrated in posterior and lateral regions around the blastopore
(Fig. 7G–J). As the blastopore constricts at later stages of epiboly,
localization becomes more prominent in cells of the blastopore
lip (Fig. 7M, N). Expression becomes less pronounced along the
posterior edge of the blastopore (Fig. 7O, R). At later stages, gsc
mRNA is seen in the cells that undergo convergent extension,
including the two terminal cells, which originally comprised the
posterior blastopore lip (Fig. 7M, N). As development continues,
gsc is seen in all cells of the developing mouth and the esophagus, and is more prominently expressed in the anterior regions of
these structures (Fig. 7Q–S). Additional regions of expression are
also noted in post-trochal right and left regions in pre-larval
specimens (Fig. 7Q–T).
During elongation stages, expression is noted in the 4d derived
left and right teloblasts (Fig. 7K, L) and in endomesodermal progeny of these cells, as well as the hindgut rudiment (Fig. 7Q–S).
Furthermore, expression is apparent in the progenitors of the
ectomesoderm (3a2, 3b2; Fig. 7R) and seen faintly in scattered
mesenchymal cells.
Hex. Hex is a divergent homeobox transcription factor involved
in anterior-posterior patterning (Thomas et al., 1998). This gene is
expressed in vertebrate anterior endomesoderm and is required for
anterior development, including the forebrain (Newman et al.,
1997; Zorn et al., 1999; Jones et al., 1999; Brickman et al., 2000;
Barbera et al., 2000; Zamparini et al. 2006). The literature also suggests a role in migrating hepatic endoderm (Bogue et al., 2000).
DEVELOPMENTAL DYNAMICS
1228 PERRY ET AL.
Fig. 9. Expression of nk2.1 during early development in C. fornicata. A–P: nk2.1 expression in embryos ranging from 24-cell–160hpf (early cleavage through late elongation stage). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right
corner of each panel (an., animal; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. A–D labeled within
each panel in A,B represent the relative position of the embryo quadrants. Animal view of nk2.1 expression in cleavage stage embryo (A) and corresponding DAPI-labeled nuclei (B). Dorsal view (C) of nk2.1 expression in the teloblasts (2ML, 2MR) and their progeny (2mL1, 2mR1) with corresponding DAPI labeling (D). Oval stage specimen (E) with corresponding DAPI labeling (F), as well as ventral view of older oval stage specimen (G)
and corresponding dorsal view (H). Various views of expression are shown from the late ovoid specimen (I–L), and also an early organogenesis
specimen (M–P). Structures are labeled following designations used in Figures 2, 3, and 5. Scale bar in P ¼ 50 mm.
Expression of hex is not observed during C. fornicata early
cleavage stages. The first detectable expression emerges within
mid to late gastrula embryos as faint labeling in a subset of cells
of the blastopore lip (Figs. 1, 8A, B; Table 1). During later stages
of gastrulation, expression is seen in the more anterior and lateral
cells of the developing mouth and esophagus, though it becomes
more diffuse at later stages (Fig. 8C–H). During organogenesis
stages expression is also noted within two deeper areas in the
head, which we presume to be neuronal cells in the vicinity of the
photoreceptors (Fig. 8E–G).
hex is also localized to the hindgut primordium during organogenesis stages (Fig. 8D). This hindgut expression persists during
later pre-larval stages (Fig. 8E–G).
Nk2.1 (ttf1). Nk2.1 is a homeodomain transcription factor
known to regulate the development of endodermal fates, but also
plays a role in the specification of the nervous system (Ciona, Ristoratore et al., 1999; Amphioxus, Venkatesh et al., 1999). Additionally, this transcription factor is reported to be a potential
downstream target of bra in the sea urchin oral ectoderm GRN
(Rast et al., 2002).
Expression is first observed in cleavage stages within each of
the four quadrants, and is localized to regions adjacent to the
nuclei (presumably the centrosomes of the macromeres, as well as
many of the micromeres, Figs. 1, 9A–D; Table 1). This localization
of transcripts is dynamic and depends on the cell cycle. Expression is no longer detected during early to mid gastrula stages.
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1229
Fig. 10. Expression of otp during early development in C. fornicata. A–L: otp expression in embryos ranging from 37–140hpf (late cleavage through
late elongation stage). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each
panel (an., animal; veg. vent., vegetal/ventral; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Animal view
of otp expression in late cleavage embryo (A) and corresponding DAPI-labeled nuclei (B), as well as vegetal ventral view of epiboly embryo (C) and corresponding DAPI nuclei (D). Embryonic quadrants A–D are labeled within each panel of A, B. Ventral view of 99hpf specimen (E) and corresponding
DAPI-labeled nuclei (F). Expression around the blastopore is observed in an epiboly specimen (G) and ovoid specimen (H). otp expression is shown
with various views for the same late ovoid specimen (I–L). Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in I, J,
and L. Structures are labeled following designations used in Figures 2 and 5. Scale bar in L ¼ 50 mm.
Towards the end of gastrulation and during the formation of the
mouth, expression appears in the anterior-most region of the stomodeum (Fig. 9E–G). Expression also expands in an anterior crescent above the mouth (Fig. 9I–K, M-O). These cells presumably
include neural precursors of the CNS (Fig. 9M–O). nk2.1 is also
expressed in what appear to be the terminal cells (Fig. 9I–K; MO). During organogenesis and pre-larval stages, expression is
noted in the shell gland (Fig. 9I, K, L, O, P). Some expression is
also seen in the anterior mouth and in the esophagus. Two more
intensely labeled lateral clusters of cells reside just outside of the
left and right sides of the developing mouth (Fig. 9I–K, M–O).
nk2.1 also continues to be expressed in the terminal cells at these
later stages (Fig. 9M–O).
During late cleavage stages, expression is observed faintly in
the progeny of the mesentoblast, 4d, including the 2ML and 2MR
teloblasts, and more prominently in two daughters (2mL1, 2mR1;
Fig. 9C, D). However during gastrulation and later stages of organogenesis, expression is no longer detected in mesentoblast progeny and no expression is detected in ectomesoderm.
Orthopedia (otp). Orthopedia is a homeobox transcription factor that is commonly associated with CNS and brain development
in the mouse. Specifically, it controls development of the hypothalamus, a derivative of the forebrain in vertebrates (Acampora
et al., 2000; Kaji and Nonogaki, 2013). In Drosophila, otp is
expressed during gastrulation in the proctodeum, and during later
stages in the CNS and posterior fates, including the hindgut and
anal plate regions (Simeone et al., 1994).
Expression of otp is first observed in the D quadrant during
cleavage stages (Figs. 1, 10A, B; Table 1). During early epiboly
stages, expression is present in cells from all four quadrants and
becomes more intense in cells along the blastopore lip (Fig. 10C–
F). At later stages, expression continues around the constricted
blastopore (Fig. 10G, H). During organogenesis stages, labeling is
observed along the anterior-lateral edges of the developing
mouth and esophagus, but expression in the posterior region of
the mouth is visibly diminished (Fig. 10J). Some anterior expression is observed in progeny of the first quartet. Elevated expression is seen in posterior ectoderm cells in the blastopore lip close
DEVELOPMENTAL DYNAMICS
1230 PERRY ET AL.
Fig. 11. Expression of six3/6 during early development in C. fornicata. A–P: six3/6 expression in embryos ranging from 120–196hpf (early elongation to late stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom
right corner of each panel (ant., anterior; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Ventral six3/6
expression is shown for a range of ovoid specimens: (A) with corresponding dorsal view in B, later ovoid (C) with corresponding dorsal view in D,
and slightly later ovoid stage (E) with a corresponding anterior view of neural precursor labeling (F). Ventral views of later ovoid stages showing
the expanded neural precursor cell labeling are presented in G, H. Various views of six3/6 expression are shown for a later ovoid specimen (I–L),
as well as multiple views from an organogenesis specimen (M–O) are presented. An anterior view of a late organogenesis specimen shows
expression around the ocelli (P). Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in I, J, L–N. Structures are
labeled following designations used in Figures 2, 3, and 5, and as follows: cb, ciliary bands; oc, ocelli; pt, prototroch. Scale bar in P ¼ 50 mm.
to the ventral midline (Fig. 10G, H). Faint expression of otp
appears along the ventral midline posterior to the blastopore (Fig.
10G, H, J). There appears to be diffuse labeling in the ectoderm at
later stages. Interestingly, an asymmetric patch of ectoderm on
the right post-trochal region expresses otp (Fig. 10I, J, L), similar
to that described for genes such as cdx, otx, and foxA. Diffuse
expression is also noted in the shell gland (Fig. 10I, K, L).
During gastrulation, otp mRNA is observed in progenitors of
ectomesoderm derived from 3a2 and 3b2 (Fig. 10G, H). Faint
expression is seen in both the endomesodermal (2mL, 2mR) and
endodermal progeny of the mesentoblast (Fig. 10E–H). During
organogenesis, otp expression is not detected in the ectomesoderm or endomesoderm, nor is it expressed in the developing
hindgut rudiment.
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1231
Fig. 12. Expression of snail2 during early development in C. fornicata. A–P: snail2 expression in embryos ranging from 130–170hpf (elongation
stages to early stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each panel (vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Ventral view of snail2
expression in ovoid stage (A) and corresponding DAPI labeled nuclei (B), as well as a later ovoid stage (C) and corresponding DAPI nuclei (D). Various views of expression are shown from a late ovoid specimen (E–H), a specimen with various views transitioning to the organogensis stage (I–L)
and various views of a specimen during organogenesis (M–P). Black arrowheads point to asymmetric post-trochal ectodermal patch of expression
in M, N, and P. Structures are labeled following designations used in Figures 2, 3, and 11. Scale bar in P ¼ 50 mm.
Six3/6. Six3/6 is a homeobox transcriptional regulator commonly associated with the development of the forebrain and
anterior neural structures including eyes or photoreceptors, and
is expressed in the chordate mouth primordium (Niimi et al.,
1999; Boorman and Shimeld, 2002; Christiaen et al., 2007;
Kumar 2009). It is also involved in the specification of the apical
plate in Platynereis (Marlow et al., 2014). Alternatively, six3/6 is
involved in the specification of the aboral (posterior) territory in
cnidarians (Sinigaglia et al., 2013).
six3/6 expression is not observed at early cleavage stages, but
appears anteriorly after the onset of gastrulation when the blastopore is beginning to constrict (Figs. 1, 11A, B; Table 1). During mid
to late gastrula stages, expression appears in additional cells
around the blastopore lip, being more prominently expressed in the
region of the anterior blastopore lip (Fig. 11C, E, G). Expression is
also noted as spots in the anterior (animal) with some faint expression observed around the anterior portion of the forming blastopore, which are presumably neural progenitors (Fig. 11A–H). Two
DEVELOPMENTAL DYNAMICS
1232 PERRY ET AL.
Fig. 13. Expression of twist during early development in C. fornicata. A–L: twist expression in embryos ranging in stages from 24-cell–170hpf
(cleavage stages to early stages of organogenesis). Please refer to the detailed descriptions in the Results section. Orientation of specimens is
located in the bottom right corner of each panel (an., animal; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield
images. A cleavage stage (A) is shown with the corresponding DAPI-labeled nuclear image (B). A–D labeled within each panel in A, B represent
the relative position of the embryo quadrants. Ovoid specimens in C and E are shown with corresponding DAPI images in D and F, respectively. A
ventral view of a late ovoid specimen (G) is shown with the corresponding dorsal view (H). Multiple views of the same organogenesis specimen
are shown in I–L. Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in E–J, L. Structures are labeled following
designations used in Figures 2 and 3. Scale bar in L ¼ 50 mm.
intense focused regions of expression persist around the animal
pole through later gastrula stages, which are presumably the forming ocelli. Very faint expression is present along the ventral midline
during mid to late gastrula stages. Expression is observed in cells of
the ciliary band, which later expands to cells of the developing
velum (Fig. 11I–K; M–O). Later stages continue to exhibit expression in the cells of the developing CNS, including the precursors of
the apical ganglia and the ocelli (Fig. 11N, P; Hejnol et al., 2007). In
these later stages, six3/6 is noted in the ventral midline cells undergoing convergent extension, which includes the terminal cells (Fig.
11E, H).
During stages of organogenesis, intense anterior signal is
observed in a symmetric pattern, which represents components
of the developing brain and the ocelli located deeper within
the developing head (Fig. 11H, J, N, P). Pre-larval specimens
show expression in the mouth, esophagus, left and right velar
rudiments, which include the developing ciliated bands, left
and right food grooves (Fig. 11I–K; Lyons et al., 2015), the
shell gland, central nervous system and neurosensory cells of
the foot (which may be the statocysts), with asymmetric
expression noted on the right side in the post-trochal ectoderm
(Fig. 11I–O).
six3/6 is detected transiently in both ectomesodermal and
endomesodermal cells during mid- to late gastrula stages. six3/6
is not detected in the developing hindgut rudiment.
EMT Markers
Snail2. Snail genes encode zinc-finger transcription factors
commonly associated with development and behavior of vertebrate neural crest. Those cells are of ectodermal origin, which
arise along the dorsal midline during closure of the neural tube
and undergo epithelial to mesenchymal transition (EMT) to adopt
a variety of fates (Baker and Bronner-Fraser, 1997a, b; Nieto,
2002). snail2 (also known as slug) is directly implicated in the
process of EMT and serves as a transcriptional repressor of Ecadherin (Knight and Shimeld, 2001). Additionally, Snail proteins
can also function as regulators of mesodermal invagination
(Hemavathy et al., 2000). Only one copy of snail has been isolated
in C. fornicata. According to phylogenetic analysis (not shown),
this clone was determined to be orthologous to vertebrate snail2.
snail2 first appears in a small population of anterior ectodermal cells during early stages of gastrulation (Figs. 1, 12A–D;
Table 1). During elongation stages, an asymmetric region of
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1233
Fig. 14. Expression of notch2 during early development in C. fornicata. A–L: notch2 expression in embryos ranging from 37-cell–170hpf (cleavage stages to early stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the
bottom right corner of each panel (an., animal; veg., vegetal; vent., ventral). DAPI labeled nuclei (blue fluorescence) accompany some brightfield
images. A–D labeled within each panel in A–D represent the relative position of the embryo quadrants. An animal view is shown of a cleavage
embryo (A) with the corresponding DAPI labeling (B). Epiboly is just beginning in the vegetal view (C) and corresponding DAPI-labeled image (D).
An early ovoid stage (E) with the corresponding DAPI labeling (F), as well as a later ovoid stage showing the ventral (G) and dorsal view (H) are
shown. Various views of an organogenesis specimen are shown in I–L. Black arrowheads point to asymmetric post-trochal ectodermal patch of
expression in G–J, L. Structures are labeled following designations used in Figures 2 and 3. Scale bar in L ¼ 50 mm.
snail2 expression begins to emerge (Fig. 12A–H) and this asymmetry persists into stages of organogenesis in an area posterior to
the blastopore on the right lateral side (Fig. 12I–P). Asymmetric
expression of another Snail family member (Pv-snail1) was also
observed in Patella (Lespinet et al., 2002). At later stages of organogenesis, expression is visible in anterior neuronal precursors
that may correspond to the apical ganglia and ocelli (Hejnol
et al., 2007; Fig. 12E–G, I–K, M–O), as well as scattered cells in
other regions of the embryo. Expression is observed in a subset of
cells around the mouth and in bilateral bands of cells that contribute to structures of the velar rim, including the food groove
and metatroch (Fig. 12I–K). In the oldest pre-larval specimens,
cells in the developing head and more specifically, the developing
CNS (ocelli and cranial ganglia) express snail2. Some of those
cells are located in the surface ectoderm and may be neuronal
precursors, whereas some are deeper cells (presumably ectomesodermal cells that have undergone EMT, Fig. 12I–P). In addition,
some cells towards the right side of the foot rudiment also
express snail2 (Fig. 12M, N).
Ectomesodermal progenitors express snail2 in mid to late gastrula stages. Though snail2 is expressed in ectomesoderm, it is
not expressed in endomesoderm. No expression of snail2 was
detected in the hindgut rudiment.
Twist. Twist is a bHLH transcription factor, which is essential
for mesoderm development (summarized by Technau and Scholz,
2003). It is involved in specification and patterning of mesoderm
and is expressed in the larval head mesoderm, as well as the stomodeum, foregut, and hindgut of polychaetes (Dill et al., 2007).
Additionally, it functions to regulate gastrulation, and is an
inducer of EMT during gastrulation events, specifically by repressing E-cadherin, which results in modulation of cell adhesion
(Gheldof and Berx, 2013; Wong et al., 2014). twist plays a key
role in regulating the development of the neural crest in vertebrates (Meulemans and Bronner-Fraser, 2004; Adams, et al.,
2008; Betancur et al., 2010).
twist is expressed transiently during early cleavage stages
(Henry et al., 2010b), and reappears in mid to late gastrula
DEVELOPMENTAL DYNAMICS
1234 PERRY ET AL.
Fig. 15. Expression of hesA during early development in C. fornicata. A–L: hesA expression in embryos ranging from 130–170hpf (elongation
stages to early stages of organogenesis). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each panel (vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Ventral views are
shown of an ovoid stage (A) with corresponding DAPI labeling (B), as well as a later ovoid stage (C) with corresponding DAPI labeling (D). Multiple
views of the same late ovoid stage embryo are shown in E–H, as well as multiple views of an early organogenesis specimen in I–L. Structures are
labeled following designations used in Figures 2 and 3, and as follows: nsc, neurosensory cell. Scale bar in L ¼ 50 mm.
embryos in the anterior and lateral edge of the blastopore (Figs.
1, 13A–F; Table 1). Faint label is also seen in the lateral ectoderm
and expands at later stages in anterior ectoderm of the head and
velum (Fig. 13E–K). twist appears to be very faintly expressed
along the posterior blastopore lip and in posterior cells including
those that have undergone convergent extension (progeny of
3c 2 and 3d 2 ; Fig. 13C–F). During stages of organogenesis,
twist is also detectable in the developing mouth and esophagus (Fig. 13G). A gradient of expression is observed in the
developing mouth, where labeling is more intense along the
anterior region and less intense along the posterior edge (Fig.
13G, J). In later stages, asymmetric expression is also noted
faintly in the right post-trochal region (Fig. 13I, J, L). twist is
expressed more intensely in bilateral bands that run along
the edge of the developing velum and include precursors of
the prototroch, metatroch, and ciliated food groove (Fig. 13I–
K). The ectoderm of the head (neuronal cells) and the proliferative zone of the developing shell also exhibit twist expression (Fig. 13H, L).
While expression is detected in the precursors of the ectomesoderm (3a2, 3b2), it is not seen in the mesenchymal progeny later
during organogenesis. twist is not detected in the endomesoderm
or in the hindgut rudiment.
Notch Signaling Members
Neurogenic locus notch homolog protein (notch2). Neurogenic
locus notch homolog protein (Notch) family members are single
pass membrane proteins that serve in contact mediated intracellular signaling and interact with their receptors Delta and
Jagged/Serrate (Lawrence et al., 2000; de Celis, 2013). Notch proteins play key roles in regulating many developmental processes
including differentiation, proliferation, and apoptosis (ArtavanisTsakonas et al., 1999). C. fornicata contains 3 variants of Notch.
For the purposes of this study, we have examined, C. fornicata
notch2, which has relevant expression during gastrulation and
particularly in the development of the digestive tract.
Expression for notch2 is seen during cleavage stages in the macromeres. (Figs. 1, 14A–D; Table 1). Further expression is observed during mid to late stages of gastrulation, appearing faintly to the sides
of the blastopore. Expression continues to spread to more anterior
cells as the embryo undergoes elongation, being excluded from the
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1235
Fig. 16. Expression of hesB during early development in C. fornicata. A–P: hesB expression in embryos ranging from 80–160hpf (epiboly stages
to late stages of elongation). Refer to the detailed descriptions in the Results section. Orientation of specimens is located in the bottom right corner of each panel (an., animal; vent., ventral). DAPI-labeled nuclei (blue fluorescence) accompany some brightfield images. Animal view of hesB
expression in epiboly stage (A) and corresponding DAPI-labeled nuclei (B), as well as an early oval/elongating stage (C) and corresponding DAPI
nuclei (D). A–D labeled within each panel in A, B represent the relative position of the embryo quadrants. Multiple views of the same later oval
stage specimen are shown in E–H, and similar multiple views are shown for a late oval specimen (I–L) and early organogenesis specimen (M–P).
Black arrowheads point to asymmetric post-trochal ectodermal patch of expression in E, F, H, I, L–N, P. Structures are labeled following designations used in Figures 2, 3, and as follows: ect, ectoderm. Scale bar in P ¼ 50 mm.
posterior lip and developing mouth, with very faint expression along
the very posterior edge (Fig. 14E–G). Late elongation stage specimens
exhibit more intense expression around the anterolateral edges of
the mouth and esophagus (Fig. 14G). Expression is also seen in the
ectoderm lateral to the mouth, and the foot rudiment (Fig. 14I–L).
Some expression is also seen in post-trochal ectoderm. Expression is
also visible in cells of the head that likely include neuronal precursors. During pre-larval stages, there is more intense expression in
rudiments of the velar lobes, in the mouth and the foot, as well as
the shell gland (Fig. 14H–L). Interestingly there is also an asymmetric
patch of ectodermal cells expressing notch2 on the right side of the
post-trochal region (Fig. 14G–J, L).
No expression of notch2 was noted in endomesoderm. However, prior to undergoing EMT, progenitors of the ectomesoderm show some faint expression of notch2 (Fig. 14E, F). At
later stages during organogenesis, expression is detectable in
hindgut endoderm, which is derived from the 4d mesentoblast
(Fig. 14I, J).
DEVELOPMENTAL DYNAMICS
1236 PERRY ET AL.
Hes. The Hes family includes basic helix-loop-helix (bHLH)
type transcription factors and its members act as repressors,
except Hes6, which is an activator protein due to its inhibitory
effect on the repressor Hes1. Hes members are downstream targets of Notch signaling (Iso et al., 2003), and are known to be
involved in neural differentiation, as well as the regulation and
maintenance of stem cells in digestive systems (Baek et al., 2006;
Kageyama et al., 2008). They are also involved in cell fate determination, and this occurs by physical interactions between cells
expressing the Notch receptor and Delta/Jagged ligands in adjacent cells. Finally, they are involved in regulating the timing of
biological events such as segmentation of the somites (Kageyama
et al., 2007). For this study, we report on the expression of two
isolated hes genes, which we have named hesA and hesB. Phylogenetic analysis (not shown) was inconclusive in determining
which of the known orthologs each C. fornicata hes gene most
closely resembles, but showed that C. fornicata HesA and B are
most closely related to the Hes1, Hes2, and Hes4 branch of Homo
sapiens. Compared to a canonical Hes sequence, we found that
both genes have the same domains as the canonical form, including its characteristic basic domain with the proline residue.
hesA expression is not observed during cleavage or early to
mid gastrula stages, but appears later during formation of the
esophagus in two bilateral clusters of cells (Figs. 1, 15A–D; Table
1). Expression is also detected in neurosensory cells located in the
foot (Fig. 15I-K; Lyons et al., 2015), which could be related to the
posterior expression seen at earlier stages (Fig. 15A–C, E-G). In
addition some expression is seen at the base of the foot rudiment
anteriorly to the left and right sides.
Expression is not seen in the progeny of the mesentoblast, but
the location of expression during later stages of gastrulation suggests that the expression may be associated with some ectomesodermal progenitors at the blastopore prior to their undergoing
EMT (Fig. 15E-H).
hesB has a more dynamic expression pattern over a broader
range of stages, and is seen first during late cleavage stages
within variable cells of all four quadrants of the embryo (Figs. 1,
16A, B; Table 1). Some faint spots of expression are noted in cells
of the anterior ectoderm (Fig. 16C, D). Mid to late gastrula stage
embryos display expression in a somewhat symmetrical pattern
flanking the blastopore (Fig. 16E–G). Expression is also noted in
cells along the anterior-lateral blastopore lip (Fig. 16I–K). Some
hesB expression is noted during stages of organogenesis in ectoderm, as well as in the right post-trochal region (Fig. 16H–P).
Increased expression is also noted along the lateral edges of the
foot rudiment and extending toward the medial area of the foot
(Fig. 16M–O).
Ectomesoderm expression is noted in the lateral blastopore
region during gastrulation stages (Fig. 16C, D). Intense expression
is observed in the latest pre-larval stages on either side of the
mouth in cells radiating outward towards the anterior and posterior regions. Some of these are ectomesodermal and others appear
to be superficial ectodermal cells. Cells expressing hesB seem to
be arranged with linear-periodic distributions (Fig. 16E–P).
Expression is noted in the velum in scattered mesenchyme cells.
Discussion
Relatively little is known about how germ layers are specified in
the Spiralia, despite the fact that it is a large branch of the Bilateria. We took advantage of newly generated EST databases in the
slipper snail, C. fornicata, and examined expression patterns for
fifteen regulatory genes known to be involved in gastrulation
and germ layer specification in other metazoans. We present
these expression patterns in the context of recently completed
fate maps for gastrulation-stage embryos (Lyons et al., 2012,
2015), which allow us to assess expression relative to specific
cells and germ layers. It is possible to compare cell-type specific
gene expression patterns between species because many spiralians share a homologous, stereotyped cleavage pattern. We discuss these findings within the framework of ongoing debates
regarding the evolution of germ layers, and the digestive tract
(Technau and Scholz, 2003; Hejnol and Martindale, 2008; Martindale and Hejnol, 2009; Lyons and Henry, 2014; Hejnol and
Martın-Duran, 2015).
Endomesoderm
ctnnb, bra, cdx, foxA, gsc, nk2.1, otp, otx, and six3/6 were all
expressed in the endomesodermal progeny of 4d (Table 1). Four
of the genes examined here also exhibited enhanced expression
within the 4d-derived teloblasts, which include bra, cdx, gsc, and
nk2.1. Two of these genes, bra and cdx, are also known to be
expressed in the paired mesoteloblasts in Patella (Lartillot et al.,
2002b; Le Gouar et al., 2003). While ctnnb, bra, foxA, and otx are
known to be involved in the highly conserved sea urchin endomesoderm GRN, in contrast, gsc, nk2.1, and six3 are part of the
sea urchin ectoderm GRN (Oliveri et al., 2008; Peter and Davidson, 2011; Li et al., 2014). We were not able to examine expression for orthologs of additional genes often found in metazoan
mesoderm, such as Gata-family genes, Blimp/Krox or Mef2,
which are either not present in our available ESTs, or have not
yet been cloned.
Our results indicate that there is conservation in terms of the
deployment of metazoan endomesodermal specification in this
representative of the Spiralia. However, while there are similarities in the particular genes that are expressed during endomesoderm specification, it is obvious that the expression domains do
not have exactly the same expression patterns, even within the
Crepidula endomesodermal lineage. Therefore, these differences
could potentially serve as a starting point for understanding
specification of sublineages in this tissue.
A Shared Toolkit for Crepidula Endo- and
Ectomesoderm
In C. fornicata, the 3a and 3b cells are bipotential precursors, and
only one daughter cell of each (the 3a2 and 3b2 cells) gives rise to
mesoderm. Given that some similar mesodermal fates differentiate
from both ectomesoderm and endomesoderm (e.g., muscle cells),
one can hypothesize that components of the endomesodermal
GRN are shared between the two. Alternatively, it is conceivable
that the genes that control ectomesoderm specification are distinct from those controlling endomesoderm specification. There
are a few examples of novel origins of mesoderm tissues that are
distinct from the eumetazoan endomesoderm, for example the
evolution of mesoderm in ctenophores (Ryan et al., 2013), and
entocodon of some cnidarian medusae (Burton, 2008). Additionally, the neural crest generates some fates in common with both
ectoderm and mesoderm (Sim~
oes-Costa and Bronner, 2015). Neural crest cells also undergo EMT like cells of the ectomesoderm in
Crepidula. In each of these cases, genes typically associated with
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1237
endomesoderm are not expressed. Thus, we were interested to
compare the expression profiles of C. fornicata 3a2, 3b2, and 4dderived lineages to examine any similarity between their repertoires of regulatory factors.
Expression in C. fornicata ectomesodermal progenitors was
noted for the following: ctnnb, bra, cdx, foxA, gsc, hesA, hesB,
notch2, otp, otx, six3/6, snail, and twist. All of those genes
except for snail2, twist, notch2, hesA, and hesB are expressed in
both ectomesoderm and endomesoderm. Most of the expression
seen in these two populations of cells was coincident over the
time intervals we examined (Figs. 1–16; Table 1). Some exceptions were found in the timing (initiation vs. termination) of certain genes such as foxA and gsc (Figs. 4 and 7; Table 1).
The expression profiles of ectomesoderm and endomesoderm
are not identical, which suggests that some aspects of their
specification and/or their cellular behaviors are different. The
spatial separation of the two sources of mesoderm has been
noted in other spiralians. For example, Lartillot et al. (2002a)
and Nederbragt et al. (2002b) questioned any homology between
ecto- and endomesoderm in the gastropod Patella. In contrast to
Crepidula, Nederbragt et al. (2002b) showed that twist, and Lartillot et al. (2002a) showed that gsc and foxA, are expressed in
ectomesoderm, but not endomesoderm. Lartillot et al. (2002a)
argued that ectomesoderm may be homologous with anterior
prechordal mesoderm of vertebrates, while posterior endomesoderm may be homologous with the tail-trunk mesoderm. An
alternative hypothesis is that differences in gene expression
might reflect different behaviors of these two forms of
mesoderm.
However, since there is some overlap in the expression of
genes in both endomesoderm and ectomesoderm, the data may
suggest that ectomesoderm is specified by a very similar set of
genes that specify endomesoderm. Given these similarities, different scenarios for the evolution of ectomesoderm can be envisioned. When ectomesoderm arose, it may have co-opted the
spiralian endomesodermal tool kit. Alternatively, ectomesoderm and endomesoderm may have had a common origin,
being separated into distinct domains (e.g., anterior vs. posterior, or ventral vs. dorsal), which subsequently underwent
changes during the course of evolution. Depending on the species, ectomesoderm can arise from different combinations of
cells from any of the four quadrants (Lyons and Henry, 2014),
and 4d has been described as an ectomesodermal cell in the
annelid, Capitella (Meyer et al., 2010a, b). Distinguishing
between these scenarios will require building GRNs and additional lineage data and lineage-specific expression of
“endomesodermal” genes in a wider range of spiralians.
Differences in Ecto- Versus Endomesodermal
Expression May Be Tied to Morphogenetic Behaviors
The differences in ecto- and endomesodermal expression may
reflect different behaviors of these two derivatives of mesoderm.
For example snail2 and twist may be necessary for the EMT that
we recently described as a key behavior of the ectomesodermal
cells that express these genes (Lyons et al., 2015). snail and twist
are necessary for EMT in the sea urchin primary mesenchyme
cells (Wu and McClay, 2007; Saunders and McClay, 2014), fly
mesoderm (Nieto, 2002), and vertebrate neural crest (Carl et al.,
1999; Hall 2000). We note that in C. fornicata, the 4d-derived
mesoderm does not undergo EMT, but is born internally (Lyons
et al., 2012; Lyons and Henry, 2014). Other species of spiralians
gastrulate by invagination, and in those cases the 4d-derived
mesoderm may become internalized by EMT (Lyons and Henry,
2014). Examining gene expression patterns in these other spiralian species would allow us to broaden our comparison of expression of snail and twist within the 4d lineage. We note that in
Patella, twist is expressed in the ectomesoderm of trochophorestaged embryo, but snail is not (Nederbragt et al., 2002b; Lespinet
et al., 2002).
Further, since all of the genes expressed in ectomesoderm
(snail2, twist, notch2, hesA, and hesB) start their expression later
compared to the expression of the other endomesodermally
expressed genes, one could argue that these five genes, and
potentially others (Figs. 12–16; Table 1), may be necessary for
ectomesoderm differentiation or morphogenesis and not necessary for its initial specification. Alternatively, ectomesoderm
specification may simply be delayed in comparison with endomesoderm. Some of these genes could have other roles in segregating cells fates within the 3a and 3b lineages. For example Notch/
Hes signaling is known to be involved in contact mediated cell
specification in other systems (Drosophila, Greenwald, 1998;
mammals, Kageyama and Ohtsuka, 1999). Likewise, twist expression has been noted later during the specification of anterior
mesoderm in brachiopods (Passamaneck et al., 2015). Given the
fact that little is known about the morphogenetic behavior of
either source of mesoderm in spiralians, it will be very interesting
to correlate gene expression and cell behavior in these animals as
more detailed descriptions of their development become
available.
Endoderm
Endoderm is derived from different blastomeres in C. fornicata
including the 4th quartet micromeres and macromeres. The
hindgut rudiment is a compact structure comprised of many
small cells derived specifically from progeny of the mesentoblast
4d (Lyons et al., 2012). We found that many of the genes examined in this study were expressed in the hindgut endoderm,
including: ctnnb, bra, cdx, foxA, gsc, hex, notch2, otp, and otx
(Tables 1 and 2). While most of these genes were expressed at all
stages examined, some were expressed relatively late (hex at
120 hpf, elongating, short oval stage; notch2 at 170 hpf, early
organogenesis), while others are turned off by 140 hpf (elongating, late oval stage; six3/6, otp).
Hindgut development in some bilaterian members is often
characterized by the expression of conserved genes, which
include bra, cdx, otp, foxA, and nk2.1 (Hejnol and Martindale,
2008; Table 2). Many metazoans display conserved hindgut
expression for bra (Kispert et al., 1994; Tagawa et al., 1998; Shoguchi et al., 1999, Woollard and Hodgkin, 2000; Arendt et al.,
2001; Croce et al., 2001; Arenas-Mena, 2013). However, unlike C.
fornicata, hindgut expression of bra was not observed for the
molluscs Patella or Haliotis (Lartillot et al., 2002b; Koop et al.,
2007). C. fornicata hindgut expression for cdx (Brooke et al.,
1998; Wu and Lengyel, 1998; Edgar et al., 2001; Le Gouar et al.,
2003; de Rosa et al., 2005; Shimizu et al., 2005; Fr€
obius and Seaver, 2006; Arnone et al., 2006), foxA (but not in nematodes or
hemichordates; Weigel et al. 1989; Arenas-Mena, 2006; Oliveri
et al., 2006; Boyle and Seaver, 2008, 2010) and nk2.1 (Venkatesh
et al., 1999; Takacs et al., 2002; Lowe et al., 2003; TessmarRaible, 2007) were also conserved compared to other metazoans.
1238 PERRY ET AL.
DEVELOPMENTAL DYNAMICS
TABLE 2. Summarized Comparison of Published Gene Expression Data Among Various Members of the Spiralia
and an Echinoderm (Sea Urchin)a
a
Each organism is noted along the left side of each column and the mode of gastrulation is listed below the organism’s name. Tissues examined are listed along the left side of each column and gene names are noted along the top of each column. Information
in the table was obtained from references cited: Patella (Lartillot et al., 2002a,b; Nederbragt et al., 2002a; Lespinet et al., 2002;
Nederbragt et al., 2002b; Le Gouar et al., 2003); Saccostrea (Kin et al., 2009); Haliotis (Degnan and Morse, 1993; Koop et al.,
2007); Platynereis (Arendt et al., 2001; de Rosa et al, 2005; Steinmetz et al., 2007; Tessmar-Raible et al., 2007; Kerner et al.,
2009; Steinmetz et al., 2010; Christodolou et al., 2010; Steinmetz et al., 2011; Pfeifer et al., 2013; Pfeifer et al., 2014; Gazave
et al., 2014; Marlow et al., 2014); Capitella (Fr€
obius and Seaver, 2006; Dill et al., 2007; Boyle and Seaver, 2008; Thamm and
Seaver, 2008; Christodolou et al., 2010; Boyle et al., 2014); Hydroides (Arenas-Mena, 2006; Arenas- Mena and Wong, 2007;
Arenas-Mena, 2013); Chaetopterus (Boyle and Seaver, 2010); Helobdella (Soto et al., 1997; Goldstein et al., 2001; Song et al.,
2004; Rivera et al., 2005); Themiste (Boyle and Seaver, 2010); Tubifex (Matsuo et al., 2005; Kitakoshi and Shimizu, 2010); Sea
urchin (Di Bernardo et al., 1999; Angerer et al., 2001; Croce et al., 2001; Minokawa et al., 2004; Oliveri et al., 2006; Walton
et al., 2006; Dunn et al., 2007; Wu and McClay, 2007; Wu et al., 2008; Wei et al., 2009; Cole et al., 2009).
In contrast to other bilaterians, gsc, hex and hesA are expressed
in hindgut tissues in C. fornicata (Figs. 7–15; Table 1). This was
surprising, as previous studies highlight the importance of gsc in
foregut tissues of arthropods (Drosophila, Goriely et al., 1996),
annelids (Platynereis, Arendt et al., 2001), molluscs (Patella, Lartillot et al., 2002a), and sea urchins (Angerer et al., 2001). Likewise, hex expression has been noted in the pharyngeal isthmus of
C. elegans (M€
orck et al., 2004).
The localization of otp to C. fornicata hindgut tissues was also
somewhat surprising since this gene is often associated with CNS
or brain development (Simeone et al., 1994; Umesono et al., 1997;
Acampora et al., 2000). However, in Patella, otp is also expressed
in the stomodaeum, as we noted here for C. fornicata (Arendt et al.,
2001; Nederbragt et al., 2002a; Steinmetz et al., 2011). Other studies, including one in the spiralian nemertean C. lacteus, reveal an
ancestral role for ctnnb in the establishment of endodermal and/or
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1239
endomesodermal fates (Schneider et al., 1996; Rocheleau et al.,
1997; Thorpe et al., 1997: Logan et al., 1999; Imai et al., 2000;
Miyawaki et al., 2003; Wikramanayake et al., 2003; Lee et al.,
2007: Momose et al. 2008; Henry et al., 2008). During earlier cleavage stages, C. fornicata GFP-labeled ctnnb mRNA becomes
restricted to the 4d endomesodermal lineage (Henry et al., 2008),
which ultimately gives rise to the hindgut. However, the localization of ctnnb to endomesodermal derivatives in other spiralians has
not been well characterized to make a sufficient comparison.
It seems somewhat surprising that we did not observe appreciable expression of endodermal genes in the macromeres and
fourth quartet micromeres during gastrulation in Crepidula. On
the other hand, with the exception of hex, notch2, otp, and otx,
message was detected in those cells during cleavage stages, where
they were generally found to be localized to regions we interpret
to be the centrosomes (see Figs. 5–16). Some of these transcripts
may become dispersed during gastrula stages. The large, yolky
cells could have also hindered visualization of those transcripts.
In addition, these cells become covered by ectoderm derived from
the animal cap during gastrulation, which also could have
obscured their visualization. Alternatively, it is possible that transcripts may have been translated, and/or the RNAs degraded in
those cells. Therefore, some of these genes may play a more general role in endodermal development, while others could be more
specific to the hindgut.
The Blastopore and Openings of the Digestive Tract
The evolution of the bilaterian gut is still debated (e.g. Martindale
and Hejnol, 2009; Martın-Duran et al., 2012; Lyons and Henry,
2014). One question relates to the origin of the mouth and anus
in the Deuterostomia, Ecdysozoa, and Spiralia, relative to the
blind gut of early-branching animals, like cnidarians, ctenophores, and acoels. Some authors argue that the mouth evolved
first and is homologous among all bilaterians (Arendt et al.,
2001; Christiaen et al., 2007; Hejnol and Martindale, 2008), while
the anus evolved later and independently in different animal lineages (Hejnol and Martın-Duran, 2015). In contrast, another
hypothesis argues that the mouth and anus evolved at the same
time through the process of amphistomy (Arendt and N€
ublerJung, 1997). These, and other hypotheses rely on comparative
gene expression analyses during the development and formation
of these structures.
In the Spiralia, the fate of the blastopore varies widely.
Depending on the species, cells of the blastopore lip can give rise
to the mouth (protostomy), the anus (deuterosomy), both mouth
and anus (amphistomy), or to neither (the blastopore may close
completely). Previous spiralian data focused on two species: the
limpet mollusc Patella, and the polychaete annelid, Platynereis
(Table 2). More recently, additional expression data have been
reported in two additional polychaetes, Hydroides and Capitella,
as well as expression data for a few genes from other annelids
and molluscs (Table 2). In Crepidula the blastopore gives rise to
the opening of the mouth (protostomy, Lyons et al., 2015). Here,
our study constitutes the most thorough examination of germ
layer markers among molluscs.
During early epiboly to flattened round stages (50–65% epiboly, 48–91 hpf), otx, foxA, bra, cdx, and otp, were expressed in
cells of the blastopore lip in C. fornicata (Figs. 1, 3–6, 10; Tables
1 and 2). otx was also observed around the entire blastopore
periphery for Patella (Nedebragt et al., 2002a) and Capitella
(Boyle et al., 2014). Likewise, blastopore expression was also
noted for FoxA1 and FoxA2 in Hydroides (Arenas-Mena, 2006),
and Patella (Lartillot et al., 2002b). Despite labeling of the entire
blastopore, enhanced posterior blastopore signal was noted for
both C. fornicata bra and cdx. Posterior localization of bra is consistent with that observed during blastopore formation in other
molluscs, including Haliotis (Koop et al., 2007), Patella (Lartillot
et al., 2002a), and annelids, Capitella (Boyle et al., 2014; also
seen in left-lateral blastopore) and Hydroides (expression noted
in left and right sides flanking posterior region of blastopore;
Arenas-Mena, 2013). Posterior expression of C. fornicata cdx has
also been observed in the mollusc Patella (Le Gouar et al., 2003),
and the annelids Capitella (Fr€
obius and Seaver, 2006) and Platynereis (de Rosa et al., 2005), but is not detected in early stages of
gastrulation for the mollusc, Gibbula varia (Samadi and Steiner,
2010). In contrast, otp expression in the blastopore was not
observed in Patella (Nederbragt et al., 2002a).
ctnnb and gsc were also expressed along the posterior blastopore during early epiboly in C. fornicata. In contrast, ctnnb is
expressed in many cells of the developing Platynereis embryo,
where varying levels of expression determine animal versus vegetal fates during embryonic development (i.e., Schneider and
Bowerman, 2007; Pruitt et al., 2014). gsc is not observed around
the blastopore for Capitella (Boyle et al., 2014), but is observed in
the 3a2 and 3b2 derived cells of the anterior blastopore and several cells along the posterior vegetal plate in Patella (Lartillot
et al., 2002b). Anterior blastopore lip expression was noted for
hex during early epiboly in C. fornicata, similar to that of
amphioxus (Yu et al., 2007).
When the blastopore begins to constrict further during later
stages in C. fornicata (late epiboly through elongating stages, 97–
140 hpf), expression patterns changed slightly, but a majority of
the genes display some expression in progeny associated with the
blastopore lip. The following genes were observed around the
entire periphery of the blastopore lip: ctnnb, otx, foxA, goosecoid,
otp, six3/6, and notch2. The posterior region of the blastopore
showed bra expression, which is similar to that noted in Haliotis
(Koop et al., 2007) and Patella (Lartillot et al., 2002a), and to the
dorsal/aboral (posterior) region in Hydroides (Arenas-Mena,
2013). cdx was observed only in the lateral regions of the blastopore in C. fornicata, which differs from the posterior blastopore
expression noted in another mollusc, Patella (Le Gouar et al.,
2003), but is consistent with the lateral regionalization of cdx following blastopore closure in the annelid, Capitella (Fr€
obius and
Seaver, 2006). Additionally, cdx was localized to a U-shaped
area, which extended around the posterior and lateral regions of
the blastopore for Platynereis (de Rosa et al., 2005). nk2.1 and
hex were observed in the anterior region of the blastopore in C.
fornicata, which includes ectomesodermal progenitors. Likewise,
nk2.1 was expressed in regions flanking the blastopore for Capitella (Boyle et al., 2014).
These data reveal that considerable variation exists between
spiralians in how genes are expressed in cells around the blastopore, and later during development in the mouth and anus. Many
genes are expressed in part or all of the blastopore lip, while very
few are exclusively expressed in the mouth, hindgut, or anus in
spiralians. The specification of these structures likely involves
other genes we have yet to examine. Unfortunately, in most species the behavior of the cells of the blastopore lip is not fully
understood, nor is it understood how those cells contribute to the
mouth, hindgut and anus. These data will be necessary before
1240 PERRY ET AL.
definitive arguments about the consecutive, or simultaneous,
evolution of the mouth and anus can be made.
DEVELOPMENTAL DYNAMICS
The Posterior Blastopore Lip
In spiralians the cells that contribute to the mouth and anus are
initially located close to one another during early development,
but those cells become separated by various morphogenetic
events that are not well understood in most species (Lyons et al.,
2015). Our recent cell lineage analysis of gastrulation stages in C.
fornicata revealed that the mouth and esophagus are made by
sublineages of each of the second and third quartet micromeres
(except the 2d lineage), which make up the blastopore lip at early
epiboly stages (Lyons et al. 2015). As the blastopore constricts,
clones of micromeres on the anterior and lateral sides largely
retain their relative positions along the anterior-posterior axis.
Cell re-arrangement largely involves some progeny becoming
displaced deeper into the blastopore, contributing to the esophagus, while the rest of the clone stays at the surface, contributing
to the anterior and lateral sides of the mouth (Fig. 1). In contrast,
we have shown that the posterior lip of the blastopore undergoes
closure via a novel zippering process (Lyons et al., 2015). Zippering is a unique example of convergence and extension, which
gives rise to an elongated assemblage of eight ciliated cells that
extend from the esophagus, anteriorly, towards the posterior end
of the embryo.
As closure of the posterior blastopore lip takes place, these ciliated cells displace progeny of 2d (more specifically 2d2) away
from the blastopore lip towards the posterior end of the embryo.
Progeny of 2d ultimately form the anus later in development.
Two cells of the posterior blastopore lip, which were located closest to the ventral midline prior to zippering (3c221, 3d221), are displaced farther towards the posterior pole and become isolated
within the clone of cells derived from 2d. Earlier investigators
confused these cells with those that make the anus, referring to
them as “anal cells” (e.g., Conklin, 1897). These cells do not give
rise to the anus in Crepidula, which forms much later in development from progeny of 2d2 (Lyons et al., 2015). Hence, we
renamed the former anal cells as “terminal cells.” Previous interpretations of gene expression in these cells (as being in the anus
in other spiralians) must be regarded with caution. Similar events
leading to closure of the posterior lip of the blastopore likely
occur in the annelids Polygordius and Platynereis, and the molluscs Ilyanassa and Patella (Woltereck, 1904; Arendt et al., 2001;
Lartillot et al., 2002a, Chan and Lambert, 2014; Lyons et al.,
2015), though they have not been formally described.
Just two cells from each of the 3c2 and 3d2 lineages remain at
the opening of the blastopore to give rise to the posterior-most
portion of the esophagus and mouth. The analysis reported here
shows that several genes are expressed in the progeny of 3c2 and
3d2 during closure of the posterior blastopore lip, including:
ctnnb, otx, foxA, bra, cdx, gsc, otp, six3/6, notch2, hes (Figs. 1–
16; Table 1). This also includes the terminal cells (bra, Fig. 5M–O,
Q–S, V; cdx, Figure 6K, L, O, P). Following gastrulation during
later development, bra and cdx continue to be expressed in these
cells along the ventral midline, with nkx2.1 also being expressed
in the terminal cells (Fig. 9G–K, M–O). Earlier localization of otp
to terminal cells in C. fornicata contradicts with many studies
that usually associate this gene with anterior fates such as the
apical organ in Patella (Nederbragt et al., 2002a) and the brain in
the planarian (Umesono et al., 1997), and annelids (Tessmar-Rai-
ble et al., 2007). However, otp is observed in hindgut tissues of
Drosophila melanogaster (Simeone et al., 1994) and posterior
ectoderm of hemichordates (Lowe et al., 2003).
Cells that are likely homologous to the two terminal cells of C.
fornicata express bra and cdx in the mollusc Patella (Lartillot et al.,
2002a; Le Gouar et al., 2003; see Lyons et al., 2015). Faint labeling
of bra was also observed in Haliotis in a posterior region judged to
be the site of the anal cells (likely the terminal cells), although this
was not linked to discrete cells of known lineage origins (Koop
et al., 2007). It is important to note that bra expression is not confined to terminal cells in these species. Whether or not bra plays a
role in the development of terminal cells is unclear; however, bra
expression may be involved in convergent extension and closure of
the posterior lip of the blastopore in those species where these
events likely take place. Others have proposed that bra plays a role
in regulating morphogenetic processes associated with convergent
extension in vertebrates and other bilaterians (Hardin, 1989;
Yamada, 1994; Conlon and Smith, 1999; Arendt, 2004). For example, although bra is expressed in endodermal cells during gastrulation in the annelid Hydroides, it is also expressed in ectodermal
tissue that eventually merges along the ventral midline in Hydroides, thereby contributing to elongation and displacement of the
mouth from the anus (Arenas-Mena, 2013). Likewise bra is
expressed in ventral cells that undergo convergent extension in the
annelid Platynereis (Steinmetz et al., 2007).
Expression of Genes in the Digestive Tract (Mouth and
Esophagus)
Many of the genes examined in this study are localized to both
the C. fornicata pre-larval/larval mouth and the esophagus (foregut), including ctnnb, otx, foxA, bra, cdx, gsc, nk2.1, otp, six3/6,
twist, and notch2 (Tables 1 and 2). Table 2 reveals that all of the
genes expressed in the mouth are expressed earlier in the blastopore. We found that nk2.1, otp, and six3/6 mark the mouth, and
not the endodermal hindgut (Table 2). nk2.1 was found to be
expressed in the anterior blastopore lip (Fig. 9E, G), while otp and
six3/6 were expressed in the posterior blastopore lip, and the posterior ectodermal ventral midline (Figs. 10G, H, 11E, H), before
being restricted to the mouth (Figs. 10J, 11J). Only otp was also
found in the 2d lineages, which gives rise to the anus, though we
did not examine embryos old enough to score otp in the anus
itself. These data suggest that the morphogenetic events that zipper closed the posterior blastopore lip, and separate the mouth
territory from the posterior end of the embryo, involve changes
in expression of regulatory factors that could be necessary for
proper mouth formation. Functional analyses of these genes will
be necessary to test this hypothesis.
A comparison of gene expression in the mouth among spiralians
(Table 2), and other bilaterians, reveals interesting similarities and
differences. C. fornicata ctnnb expression is comparable to that
observed in the ectoderm around the stomodaeum of the mollusc,
Haliotis (Koop et al., 2007). Likewise, bra is also observed in the
foregut of annelids (Arendt et al, 2001; Boyle et al., 2014), molluscs
(Lartillot et al., 2002a; Koop et al, 2007), echinoderms (Shoguchi
et al., 1999; Croce et al., 2001) and hemichordates (Tagawa et al.,
1998). However, some localization of C. fornicata cdx in foregut
tissue contrasts with more highly conserved hindgut expression
across the Metazoa, though cdx is also expressed in the hindgut of
C. fornicata (Brooke et al., 1998; Wu and Lengyel, 1998; Hinman
et al., 2000; Edgar et al., 2001; Le Gouar et al., 2003; de Rosa et al.,
DEVELOPMENTAL DYNAMICS
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1241
2005; Fr€
obius and Seaver, 2005; Shimizu et al., 2005; Arnone
et al., 2006). Foregut expression for foxA, gsc and nk2.1 appears to
be largely conserved when compared to other spiralians and bilaterians (Table 2). otp expression in C. fornicata foregut tissues differs from conserved patterns, where it is usually associated with
brain and sensory cells among arthropods (Simeone et al., 1994),
annelids (Tessmar-Raible et al., 2007), molluscs (Nederbragt et al.,
2002a) and platyhelminthes (Umesono et al., 1997); but anterior
neural expression is also seen in C. fornicata, as well (Fig. 10G, H,
J). Like C. fornicata, ectoderm around the stomodaeum is similarly
noted for otx expression in Patella (Nederbragt et al., 2002a) and
Platynereis (Arendt et al., 2001; Steinmetz et al., 2011). Expression
of six3/6 is commonly associated with chordate mouth formation
(Christiaen et al., 2007), and is similarly observed in the mouth and
esophagus of C. fornicata. twist is associated with foregut specification in Capitella (Dill et al., 2007), which is similar to the mouth
and esophagus expression observed in C. fornicata. Expression of
snail2 is associated with the pre-larval mouth and not the esophagus of C. fornicata. Snail2 is present in the foregut epithelium in
Capitella (Dill et al., 2007), but not in the mollusc Patella (Nederbragt et al., 2002b). hesA and hex are localized to the C. fornicata
esophagus. hes is involved with oral ectoderm specification in the
sea urchin (Minokawa et al., 2004), in the planarian stomodaeum
(Gazave al., 2014), but not in the leech, Helobdella, where it is in
the posterior addition zone (Song et al., 2004). hex expression is
found in the prospective dorsal endoderm (foregut) of the hemichordate S. kowalevskii (Lowe et al., 2006).
Overall, these data suggest that comparing mouth development
between different spiralians may be a fruitful area of future
research, as the relationship of the blastopore and the mouth, and
the patterns of gene expression in these domains vary between
species. If the bilaterian mouth is indeed homologous, then the variation seen can be used to understand what aspects of mouth formation are labile during evolution, and which are resistant to
change. These differences could be tied to variations in the process
of gastrulation (Table 2), or in the clonal contributions to the cells
of the mouth and foregut. For instance, some of the genes examined were expressed in the entire circumference of the mouth (e.g.,
bra, cdx; Figs. 5 and 6), while others were more restricted to the
anterior (six3/6, hesA; Figs. 11 and 15), or lateral (hex; Fig. 8) sides.
These different expression domains appear to correlate with clonal
lineages, and may reflect differences in cellular structure or function that should be examined in future studies.
Conserved Expression of Genes in the Digestive Tract
(Anus)
Lineage tracing in C. fornicata has demonstrated that the
endodermal hindgut arises from several daughter cells of the
4d lineage (Lyons et al., 2012), while the ectodermal anus is
derived from the 2d2 lineage (Lyons et al., 2015). At 12 days
post fertilization, the opening of the anus forms as it fuses
with the endodermal hindgut. The current gene expression
analysis stops before the definitive anus forms. While we did
not score for expression in the anus itself, we were able to
score for expression in the hindgut rudiment, terminal cells,
and the posterior 2d clone, which eventually forms the anus in
Crepidula.
Many of the genes expressed in hindgut endoderm were also
expressed in the overlying 2d clone during late epiboly and early
elongation stages, but at later stages, ventral midline staining in
the ectoderm (derived from 2d, 3c, and 3d) is quite reduced. At
later stages, we were only able to see expression of otx in the 2d
clone where the anus will eventually form (Fig. 3I–K, M–O). Thus,
while the ectodermal and endodermal components of the terminal
portion of the digestive tract initially express many of the same
genes, it is likely that the ectodermal lineages that make the anus
and other nearby ectodermal cells (e.g., TCs), express only a subset of these genes.
While the bilaterian mouth has been argued to be homologous
(Arendt et al., 2001; Hejnol and Martindale, 2008), the anus may
have evolved independently on multiple occasions (Hejnol and
Martindale, 2008; Hejnol and Martın-Duran, 2015). Hejnol and
Martindale (2008) argued that when an anus evolves it might coopt components of the endodermal hindgut GRN. In fact, most of
the genes that are reported to be expressed in the anus in spiralians are also expressed in the hindgut. The ability to compare
gene expression in the context of homologous cell lineages
makes the spiralians particularly useful. For example, lineage
tracing in C. fornicata revealed that the ectodermal anus arises
from 2d2. In Capitella both anus and hindgut arise from the 4d
lineage as a common ectodermal structure, and so it may not be
surprising that many of the genes expressed in the anus are also
expressed in the hindgut (Meyer et al., 2010b).
Conclusion
This study lays the groundwork for building GRNs during the
process of germ layer specification in C. fornicata. If we are to
tease apart how the gene regulatory networks controlling mouth
and anus formation evolved, we must understand how gastrulation takes place and how specific cell lineages contribute to these
structures. While some regard regulatory factors as “markers” for
cell fate specification, they also play roles in controlling morphogenetic events. Given that spiralians exhibit a wide range of gastrulation modes (Table 2; Lyons and Henry, 2014; Arenas-Mena,
2013), they present an opportunity to assess the role of specification versus cell behavior in the evolution of gastrulation, germ
layer formation (e.g. mesoderm) and gut development.
Experimental Procedures
Animal Care and Handling
Adult C. fornicata were harvested from local waters near Woods
Hole, MA, by the Marine Resources Center at the Marine Biological Laboratory. Embryos were collected and reared, as previously
described (Henry et al., 2006, 2010a, 2010b; Lyons et al., 2012,
2015).
Fixation and Histology
Embryos and larvae were fixed for one hour at room temperature
in a 3.7% solution of ultrapure formaldehyde (Ted Pella, Inc.,
Redding, CA) dissolved in filtered sea water (FSW), with added
Instant Ocean Aquarium Sea Salt Mixture (United Pet Group,
Blacksburg, VA), as previously described (Lyons et al., 2015). Following fixation, embryos were rinsed with three sterile 1X PBS
washes (1XPBS:1.86mM NaH2PO4, 8.41mM Na2HPO4, 175mN
NaCl, pH 7.4), followed by three washes in 100% methanol.
Embryos were stored in 100% methanol at -80degC. Following in
situ hybridization (see below), embryos were incubated in a solution of 0.5 mg/ml DAPI (Life Technologies, Grand Island, NY)
1242 PERRY ET AL.
TABLE 3.
DEVELOPMENTAL DYNAMICS
Accession
List of Primers Used to Prepare ISH Probesa
Gene
Forward primer
Reverse primer
HM040900
HM040889
KM434315
KM434317
HM046939
KP902717
ctnnb
bra
cdx
foxA
gsc
hesA
KP902718
hesB
KP885704
KM434318
KM434321
KM434316
KM434320
KP885705
KP885706
HM040896
hex
nk2.1
notch2
otp
otx
six3/6
snail2
twist
CGATAGATGAGCAGTTATCAGATG
GAGAGAAAGTGGACCCTGATGCCCAG
CAGCCATGGAGACAGCCCAGTAC
CTATGTCCTCCATGGGGTCCATGG
GAGATGGTCCGTAGCGGTCTG
CCGACAAGTACAGGCTTGGCTTC outer
GGCTTGGCTTCAATGAGTGCGCC:nest
CGAGAATCAACAGCTGTCTGTCG outer
GCTGTCTGTCGCAGCTCAAGTCC:nest
GCTGTCGTCGTCAGGCATGTTTAAAC
CATGTCCCTCAGTCCCAAACAC
GAGGAACTCATCACGGCTGAG
GGTCTGGAGATGGACGAGG
CACTAACACCACCACGGGAAGC
GAAAGCGGGGACATTGAGCGTC
CCCTCCACATCGTCCACACC
ATGGTGCTGGAGCGGCAGACA
GGTCTGTGTCAAACCATATG
GAGCCCTCTGCCCTGTCCGTC
GGAACTCCTTCTCCAGCTCCAGTC
GAGTCGCCTGCACATCATGAGG
CTTGAACCAGACCTCGACTCTC
CTGACTTCACTAGACTTGGCCTC:outer
GACTTGGCCTCACTCGTCACTGT: nest
CAAGGGGCTGTAGACGGACGAAT outer
GACGGACGAATCAAACGACATTT:nest
CAGAGTCCGGTATGATGTCGTCGTC
GTCTGGGGAATCCTTGGTGTCC
GACCCCACATCCCCGAGCC
CTATTTGACCGCTGCTGC
CTCAAAGGCTCTGGAACTCATTCATC
CTCTCTGGCGCCGGTTCTTG
TACCTCACCGGCTCGTGCTC
CACTGGCCGGCCATGGAGCC
Probe
length
1913bp
650bp
582bp
978bp
290bp
966bp
926bp
566bp
732bp
1200bp
330bp
740bp
515bp
1500bp
877bp
a
The NCBI accession number is presented along the far left column for each of the clones examined in this study. Gene-specific
forward and reverse primers used to isolate each C. fornicata clone are listed next in 50 - to 30 - orientation. For each clone used
for ISH, the DIG-labeled probe length is listed in the far right column.
dissolved in 1X PBS. DAPI incubation lasted for 10 min in the
dark, followed by 3 washes in 1X PBS/0.1% Tween. Specimens
were stored at 4degC in 80% glycerin/20% 1X PBS until imaged
(see below).
cDNA Library Preparation and Analysis
Three different C. fornicata EST databases were used to identify
and clone various genes known to be involved in germ layer
specification, gastrulation and development of the digestive tract
(Lengyel and Iwaki, 2002; Technau and Scholz, 2003; Heath,
2010; Hejnol and Martın-Duran, 2015). All the developmental
material used to prepare these libraries came from animals collected at the Marine Biological Laboratory in Woods Hole, MA.
These include a previously published C. fornicata 454 EST library
prepared by Henry et al. (2010a), and two new libraries described
below.
A second EST library was prepared in the lab of Dr. Cristina
Grande (Centro de Biologıa Molecular Severo-Ochoa, UAM-CSIC,
Madrid, Spain). The RNA used to prepare this library was
obtained from early cleavage stage embryos up through advanced
veliger larval stages (two to four weeks of age, but prior to hatching). These embryos and larvae were collected and stored in RNA
later at -80degC. Total RNA was prepared using Trizol following
the manufacturer’s instructions (Life Technologies). The embryos
and larvae were homogenized using sterile plastic pestles. To prepare the cDNA, approximately 3 mg of extracted RNA was combined from each stage collected. The RNAseq library was
constructed using TruSeq RNA Sample Prep Kit v2 (Illumina, San
Diego, CAA) following the manufacturer’s instructions. The RNAseq library was quantified using the Qubit HS DNA Assay and
sequenced in 3 HiSeq 2000 PE100 lanes (Illumina). Assembly was
done with Trinity assembler software (Grabherr et al. 2011). After
assembly, contigs less than 200 bp in length were removed from
our dataset. Assessments of the quality of read data were performed using FastQC (available at: http://www.bioinformatics.
bbsrc.ac.uk/projects/fastqc/) and the statistics on the final
trimmed assembly are as follows: Trinity transcript reads (contigs) ¼ 326,118, mean read (contig) length ¼ 564 bp, N50 read
(contig) length ¼ 717 bp; minimum read (contig) length: 201;
maximum read (contig) length: 28,997; median read (contig)
length: 350; number of reads (contigs) >¼1kb: 37,979; number
of bases in all reads (contigs): 183,902,140.
Finally, a third set of EST libraries was prepared in the lab of
Dr. Joel Smith (Marine Biological Laboratory, Woods Hole, MA).
A total of 24 Illumina RNAseq libraries from various embryonic
stages were prepared and the sequence data was pooled prior to
assembly. The 18 libraries were collected from a single brood at
consecutive time points: 147 to 223 embryos were collected in
Trizol approximately every 90 min starting at 11.5 hpf (hours
post fertilization; approximately 4 to 8-cell stage) to 40 hpf
(cleavage stages, pre 4th quartet). The 6 remaining libraries were
collected from a different brood at 22 hpf after treatment with
either U0126 (a MAPK inhibitor) or DMSO at the 4-cell stage.
These samples contained 115 to 139 embryos each. Total RNA
extraction from Trizol and Ribo-Zero low input (Illumina) was
performed as recommended by the manufacturer’s instructions.
The RNA quality before and after the removal of ribosomal RNAs
was assessed on a Bioanalyzer using RNA pico chips (Agilent
Technologies, Santa Clara, CAA). The RNAseq libraries were constructed using ScriptSeq (Illumina), with a 60-sec fragmentation
time, and otherwise following the manufacturer’s instructions.
ERCC control RNA (available from Invitrogen/Life Technologies)
was added before library preparation. The libraries were size
selected with a 2% Pippin prep gel (Sage Science, Beverly, MAA)
for 350–600 bp. The quality of the libraries before and after the
REGULATORY FACTORS IN THE MOLLUSC Crepidula 1243
size selection was assessed on a Bioanalyzer using DNA high sensitivity chips (Agilent). Bi-directional sequencing was done using
the Illumina HighSeq 1000 platform, and the assembly of this
combined library was done with Trinity assembler software
(Grabherr et al. 2011): Trinity transcript reads (contigs) ¼
285,503, mean read (contig) length ¼ 712 bp, N50 read (contig)
length ¼ 1145 bp; minimum read (contig) length: 201; maximum
read (contig) length: 30,863; median read (contig) length: 389;
number of reads (contigs) >¼1kb: 71,337; number of bases in all
reads (contigs): 203,168,019.
DEVELOPMENTAL DYNAMICS
Sequence Analysis
Assembled consensus sequence files from each EST database
were uploaded to the software program Geneious (Auckland, New
Zealand). Using publicly available invertebrate homologous protein sequences from NCBI or UniProtKB, tBlastn alignments
against each C. fornicata EST database were performed individually for each of the genes of interest in the Geneious program.
Any C. fornicata EST that aligned with a gene of interest was
then submitted to BlastX (NCBI) to verify the identity of the EST.
Each identified C. fornicata nucleotide sequence was also used as
a query sequence for additional Blastn alignments in Geneious
against the C. fornicata ESTs to further extend the nucleotide
sequence in the 50 - or 30 -direction. Extended sequence information was compiled and assembled using Sequencher software
(Gene Codes Corporation, Ann Arbor, MI) and consensus sequences verified again for identity against the NCBI database using
BlastX. Only one discrete version of each gene was found in the
EST datasets, with the exception of notch (three copies), twist
(two copies), and hes (two copies). Phylogenetic analysis was
conducted for snail2, hesA and hesB sequences (data not shown).
snail2 was analyzed and confirmed to be orthologous to the vertebrate Snail2, which is reflected in the naming scheme, even
though snail1 has not been identified in the available C. fornicata
EST collections. hesA and hesB grouped within the vertebrate
Hes1, 2 and 4 branches, but neither could be definitively
grouped. Therefore, we chose to name these C. fornicata genes
hesA and hesB, as to avoid confusion in terms of orthology.
Cloning
Using specimens stored in RNAlater (Life Technologies), C. fornicata RNA was isolated from both gastrulating embryos and preveliger larvae. High quality total RNA was extracted using TriZol,
and purification methods followed the manufacturer’s suggested
protocol. The purity and concentration of total RNA was verified
with a NanoDrop ND-1000 Spectrophotometer (Thermo Scientific, Wilmington, DE) and approximately 1mg of total RNA from
each developmental stage (epiboly and pre-veliger stages) was
used to synthesize cDNA (iScript cDNA Synthesis kit, Bio-Rad,
Hercules, CA).
Gene-specific primers were designed for each gene of interest
and those can be found in Table 3. Due to the presence of GCrich regions, PCR amplification reactions were performed with Q5
High Fidelity DNA polymerase and Q5 High GC enhancer buffer
(New England Biolabs, Ipswich, MA), according to manufacturer
suggested ratios. Amplified PCR products were run on 1% agarose gels, gel purified (GeneClean Turbo kit, MP Biomedicals,
Solon, OH) and cloned into pGem-T Easy vector (Promega, Madison, WI). All clones were verified by sequencing at the University
of Illinois’ Carver Biotechnology Center (Urbana, IL) and sequences assembled using Sequencher software (Gene Codes Corp.,
Ann Arbor, MI).
Gene Expression Analyses
Gene specific DIG-labeled probes ranged from 290 nt to 1.2 kb in
length, depending on available sequence data (see Table 3). Linearized template DNA (amplified from plasmid DNA with T7/SP6
primers) was used to synthesize RNA probes with either T7 or SP6
RNA polymerase (Life Technologies) and DIG-labeling mix
(Roche, Indianapolis, IN). Reactions were purified with RNeasy
MinElute Cleanup kit (Qiagen, Valencia, CA) and probe concentrations were verified on a NanoDrop ND-1000 Spectrophotometer. The in situ hybridization protocol was modified from
Finnerty et al. (2003) and is similar to that described by Henry
et al. (2010b), but with the following modifications. The proteinase K digestion was eliminated and following rehydration in PTw
(1X PBS, 0.5% Tween 20), embryos were washed in two acetic
anhydride washes. Hybridizations were conducted overnight at
61degC with a probe concentration of 1 ng/ml followed by graduated washes into 2X SSC at 61degC, 0.2X SSC at 61degC, and
finally PTw at room temperature. The alkaline phosphatase (AP)
buffer was modified slightly to eliminate salt precipitation (5 mM
MgCl2, 100 mM NaCl, 100 mM Tris pH 9.5, 0.5% Tween 20) and
NBT/BCIP was used for probe visualization, by incubation at
room temperature in the dark. Following in situ hybridization,
specimens were incubated in Hoecsht 33342 (Life Technologies,
1:10,000 dilution in PTw) for 10 min at room temperature for visualization of nuclei, followed by three washes in 1X PBS.
Microscopy
Fixed embryos processed for in situ hybridization, were mounted
on Rain-X-coated (ITW Global Brands, Houston, TX) glass slides
in 80% glycerol/20% 1X PBS. Coverslips were prepared as
described in Lyons et al. (2015). Specimens were visualized on a
Zeiss Axioplan microscope (Carl Zeiss Inc., Munich, Germany)
and a Spot Flex camera (Spot Imaging Solutions, Sterling
Heights, Michigan) was used for imaging. Multifocal stacks of
brightfield images were combined and flattened using Helicon
Focus stacking software (Helicon Soft Ltd., Kharkov, Ukraine).
Acknowledgments
The authors thank the Marine Biological Laboratory and especially
the directors of the Embryology course, Drs. Richard Behringer and
Alejandro Sanchez Alvarado for supporting this research. We
thank Dr. Joel Smith for his support in the preparation of one of
the C. fornicata EST databases. We also thank Dr. Nathan Kenny
(School of Life Sciences, Chinese University of Hong Kong, Shatin
Hong Kong) for his contribution in preparing the data used from
the C. fornicata EST library prepared in Spain. DCL thanks Dave
McClay for his support. C.G. is currently a “Ramon y Cajal” postdoctoral fellow supported by the Spanish MICINN and the UAM,
and funded by project CGL2011-29916 (MICINN). This research
was supported by NSF grant 1121268 to Dr. J.Q.H. (JJH).
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